diff --git a/NEXT_TASKS.md b/NEXT_TASKS.md index 66379c3a..b660182c 100644 --- a/NEXT_TASKS.md +++ b/NEXT_TASKS.md @@ -360,19 +360,56 @@ errors, e.g. sulfite → CHEBI:16731 *(E)-cinnamaldehyde* instead of CHEBI:17359 the primary full text isn't retrievable, keep edges to abstract-supported/directly-implied claims and file the rest as a KNOWLEDGE_GAP (000176 is the worked example). -**PENDING — enrich the single-edge records via a full causal-graph run.** Six 2-node -records already carry one hand-authored `downstream` edge (donor→acceptor) from original -curation but have not had an Edison causal-graph pass: the DIET trio -`Geobacter_Clostridium_DIET` (000031, PMID:28287150), -`Geobacter_Methanosaeta_DIET` (000032, doi:10.1039/C3EE42189A), -`Geobacter_Methanosarcina_DIET` (000033, PMID:24837373); and the syntrophies -`Syntrophobacter_Methanobacterium_Syntrophy` (000068), `Syntrophobacter_Methanospirillum_Syntrophy` -(000069), `Syntrophomonas_Methanospirillum_Syntrophy` (000070). A full run could add -mechanistic nodes/edges (e.g. conductive pili / OmcS-OmcZ cytochromes, conductive-material -mediation, formate-vs-H2 routes, reverse/feedback edges) beyond the single existing edge. -Run `just research-community-causal CommunityMech:0000NN` per record, then curate -conservatively as usual. NB: stray untracked `*.yaml.bak` backups exist alongside these in -`kb/communities/` (gitignored, not in the repo) — target the `.yaml` files. +**DONE — single-edge record enrichment (#254).** All six 2-node records got an Edison +causal-graph pass and conservative curation. What actually landed, per record: + +- **000070 `Syntrophomonas_Methanospirillum`** — reverse feedback edge (methanogen H2 + scavenging → enables butyrate β-oxidation). Best evidence was already in the cached + abstract, not in the Edison report: PMID:16345745 reports both the dependency + ("Growth and degradation of fatty acids occur only in syntrophic association with + H(2)-using bacteria") and the perturbation ("The addition of H(2) … stopped growth and + butyrate degradation"). No new reference needed. +- **000069 `Syntrophobacter_Methanospirillum`** — reverse feedback edge, on the + axenic-vs-syntrophic contrast in PMID:9828440 (exact pair: M. hungateii). +- **000068 `Syntrophobacter_Methanobacterium`** — reverse feedback edge + **fixed a real + misattribution**: two evidence items quoted the Harmsen M. *hungateii* passage but were + explained as establishing M. *formicicum* (this record's partner). Downgraded to PARTIAL, + added exact-pair support from PMID:29611893, and filed discussion + `kg-syntrophobacter-methanobacterium-partner-attribution`. +- **000033 `Geobacter_Methanosarcina`** — new `T6SS-Associated Delay of DIET Establishment` + interaction + NEGATIVE edge to the DIET node (>30 d lag wild-type vs very little lag for + the Hcp-deficient mutant, PMID:37650614, OA full text cached). Curated as *T6SS-associated*, + not T6SS-caused: the mutant is pleiotropic (also reduces Fe(III) oxide faster). +- **000031 `Geobacter_Clostridium`** — no new edge; filed CONTROVERSY discussion + `kg-geobacter-clostridium-contact-dependence-contested`. **This record's core framing is + contested**: the follow-up study (PMID:34939136) concludes the interaction is *mediated* + (putative cobamide), and its full text reports that 0.22-µm-filtered cell-free spent + medium reproduces the metabolic shift — which would make pili contact unnecessary. + Curator decision still open on whether to re-scope the record away from `community_category: + DIET`; not applied unilaterally. +- **000032 `Geobacter_Methanosaeta`** — no new edge (000176 precedent). Edison proposed an + acetate cross-feeding node, but its quotations came from secondary reviews; the primary + (doi:10.1039/C3EE42189A) is cached abstract-only and is silent on acetate. Filed as + `kg-geobacter-methanosaeta-acetate-route-unresolved`. + +Verification: all 6 `just validate` clean; snippet audit MATCH 4086→4101 (all 12 new +snippets match, zero new mismatches) and caching PMID:29611893 OA full text also cleared the +3 pre-existing 000068 Methods-snippet mismatches (169→166 repo-wide); network-integrity audit +clean for all 6. New reference caches: PMID:34939136, PMID:37650614. + +**Follow-ups this batch surfaced:** +1. **000031 re-scoping decision** (above) — the highest-value open item. +2. **Li et al. 2024** (`doi:10.3390/w16243551`, *Water*) has exact-pair graded-formate + perturbation data for 000068 (5–10 mM promotes, ≥30 mM inhibits; FDH/hydrogenase + transcript downregulation). **Not ingested**: the journal is not in PubMed and has no PMC + record, so `scripts/cache_fulltext.py` can't verify its snippets. Would need a + DOI-based full-text cache path. +3. `just validate-references` is a **no-op** — it reports `Total checks: 0` even on untouched + files. `scripts/evidence_snippet_audit.py` is what actually validates snippets. Worth + fixing or documenting, since the justfile recipe implies coverage it isn't providing. + +NB: stray untracked `*.yaml.bak` backups still exist alongside these in `kb/communities/` +(gitignored, not in the repo). **Edison auth (resolved 2026-07-21):** the key was refreshed in `.env` (`EDISON_API_KEY`) and authenticates (HTTP 200). The stale-key shadowing footgun is diff --git a/kb/communities/Geobacter_Clostridium_DIET.yaml b/kb/communities/Geobacter_Clostridium_DIET.yaml index 7c822871..278f61e6 100644 --- a/kb/communities/Geobacter_Clostridium_DIET.yaml +++ b/kb/communities/Geobacter_Clostridium_DIET.yaml @@ -498,3 +498,55 @@ related_ingredients: metals_present: [] metal_relevance: SIGNIFICANT metal_notes: Metal/REE detected via environmental factor measurements +discussions: +- discussion_id: kg-geobacter-clostridium-contact-dependence-contested + prompt: > + Is the G. sulfurreducens-induced metabolic shift in C. pasteurianum actually caused by + contact-dependent direct interspecies electron transfer along conductive pili, or by a + diffusible mediator (candidate: a cobamide) released by G. sulfurreducens? + kind: CONTROVERSY + status: OPEN + attaches_to: + - ecological_interactions#Acetate Oxidation and Direct Electron Transfer + - ecological_interactions#Glycerol Fermentation with DIET-Induced Metabolic Shift + rationale: > + This record asserts contact-dependent DIET through electrically conductive pili + (nanowires) as the established mechanism, following the original coculture study + (PMID:28287150), which inferred interspecies electron transfer rather than + demonstrating contact dependence. A follow-up study by the same group + (PMID:34939136) set out to identify the mechanism and concluded that the interaction + is MEDIATED: it proposes that G. sulfurreducens releases cobamide molecules that act + on the C. pasteurianum glycerol dehydratase, with electron entry via a transmembrane + flavin-bound polyferredoxin / cytochrome b5-rubredoxin route offered only as a + putative reinforcement. The published full text (not open access, so only the + abstract is snippet-verifiable here) additionally reports that 0.22-um-filtered, + cell-free G. sulfurreducens spent medium reproduces the metabolic shift, which would + make Geobacter cells - and therefore pili contact - unnecessary for the shift. + Until that contrast is verified against the full text, the nanowire/contact-dependent + framing in this record's description, its `Acetate Oxidation and Direct Electron + Transfer` interaction, and its DIET `community_category` should be treated as + CONTESTED rather than established. Resolving this may require re-scoping the record + away from DIET; that is a curator decision, not applied here. + evidence: + - reference: PMID:34939136 + supports: PARTIAL + evidence_source: IN_VITRO + snippet: It was assumed that this metabolic shift of the fermentative species resulted + from an interspecies electron transfer + explanation: The follow-up study characterizes the interspecies-electron-transfer reading + of the original experiment as an assumption, not a demonstrated mechanism. + - reference: PMID:34939136 + supports: REFUTE + evidence_source: IN_VITRO + snippet: 'C. pasteurianum-G. sulfurreducens interaction inducing a metabolic shift is + mediated' + explanation: Key-point conclusion that the interaction is mediated (via a diffusible + molecule), contradicting the contact-dependent nanowire mechanism asserted by this record. + - reference: PMID:34939136 + supports: SUPPORT + evidence_source: IN_VITRO + snippet: 'a putative interaction model was proposed: G. sulfurreducens produces cobamide + molecules that possibly modify C. pasteurianum metabolic pathway at the key enzyme + glycerol dehydratase' + explanation: States the competing diffusible-cobamide mechanism, and marks it as putative - + so it must not be asserted as an interaction in place of the current one. diff --git a/kb/communities/Geobacter_Methanosaeta_DIET.yaml b/kb/communities/Geobacter_Methanosaeta_DIET.yaml index 9dbad3ae..a054f654 100644 --- a/kb/communities/Geobacter_Methanosaeta_DIET.yaml +++ b/kb/communities/Geobacter_Methanosaeta_DIET.yaml @@ -372,3 +372,37 @@ metals_present: [] metal_relevance: SIGNIFICANT metal_notes: Metal/REE detected via environmental factor measurements; Metal/REE detected via keyword matching in description (context-validated) +discussions: +- discussion_id: kg-geobacter-methanosaeta-acetate-route-unresolved + prompt: > + Alongside DIET-driven CO2 reduction, does M. harundinacea also cross-feed on the + acetate generated by G. metallireducens ethanol oxidation, and what fraction of + methane comes from each route in this defined coculture? + kind: KNOWLEDGE_GAP + status: OPEN + attaches_to: + - ecological_interactions#Ethanol Oxidation and Direct Electron Transfer + - ecological_interactions#Direct Electron Acceptance and Methanogenesis + rationale: > + A causal-graph pass on this record proposed an additional acetate cross-feeding node + and an acetate -> methanogenesis edge, on the reasoning that Methanosaeta is + classically an acetoclastic genus and that ethanol oxidation by G. metallireducens + yields acetate, giving an overall stoichiometry near 1.5 mol CH4 per mol ethanol. + That claim is NOT curated here as an interaction: the primary source + (doi:10.1039/C3EE42189A) is cached abstract-only and its full text was not + retrievable, and the abstract states only that M. harundinacea "accepted electrons + via DIET for the reduction of carbon dioxide to methane" - it does not report acetate + cross-feeding, the acetate/DIET split, or the per-route methane stoichiometry. The + supporting quotations offered for the acetate route came from secondary reviews + attributing findings to this study, which per this repo's conservative-curation rule + (see CommunityMech:000176) is not a sufficient basis for an exact-system causal edge. + Resolving this needs the primary full text or a radiotracer/isotope partition + experiment in this exact coculture. + evidence: + - reference: doi:10.1039/C3EE42189A + supports: PARTIAL + evidence_source: IN_VITRO + snippet: Transcriptomic, radiotracer, and genetic analysis demonstrated that M. harundinacea + accepted electrons via DIET for the reduction of carbon dioxide to methane + explanation: The abstract demonstrates only the DIET/CO2-reduction route; it is silent on an + acetate cross-feeding route, which is why that edge is filed as a gap rather than curated. diff --git a/kb/communities/Geobacter_Methanosarcina_DIET.yaml b/kb/communities/Geobacter_Methanosarcina_DIET.yaml index d8456382..94b9dbc5 100644 --- a/kb/communities/Geobacter_Methanosarcina_DIET.yaml +++ b/kb/communities/Geobacter_Methanosarcina_DIET.yaml @@ -201,6 +201,66 @@ ecological_interactions: evidence_source: IN_VITRO snippet: Direct interspecies electron transfer (DIET) is potentially an effective form of syntrophy in methanogenic communities, but little is known about the +- name: T6SS-Associated Delay of DIET Establishment + description: 'Geobacter metallireducens expresses a type VI secretion system (T6SS) during + the initiation of DIET cocultures, and this is associated with a long delay before the + partnership becomes productive. Cocultures started with wild-type G. metallireducens showed + a lag of more than 30 days before methane production, whereas cocultures started with an + Hcp-deficient (T6SS structural protein) mutant had very little lag and established DIET + with M. barkeri faster. The sign of this edge is NEGATIVE: the wild-type T6SS-expressing + state suppresses the rate at which the DIET partnership is established. The molecular + attribution is confounded and must not be curated as demonstrated antagonism toward the + archaeon - the same mutant also reduced Fe(III) oxide faster than wild type, a phenotype + not expected from loss of the T6SS, and the authors state that the effect of T6SSs on + archaeal cells still requires study. Encoded as a delay of establishment, not as + predation, killing, or toxin delivery to M. barkeri. + + ' + interaction_type: COMPETITION + source_taxon: + preferred_term: Geobacter metallireducens + term: + id: NCBITaxon:28232 + label: Geobacter metallireducens + biological_processes: + - preferred_term: protein secretion by the type VI secretion system + term: + id: GO:0033103 + label: protein secretion by the type VI secretion system + downstream: + - target: Ethanol Oxidation and Direct Electron Transfer + description: NEGATIVE edge - the wild-type T6SS-expressing state of G. metallireducens + delays establishment of the DIET partnership (>30 d lag vs very little lag for the + Hcp-deficient mutant). Demonstrated by knockout phenotype in this exact coculture, but + the mutant is pleiotropic, so the causal attribution to the T6SS specifically is + HYPOTHESIZED rather than proven. + evidence: + - reference: PMID:37650614 + supports: SUPPORT + evidence_source: IN_VITRO + snippet: there was a lag period of more than 30 d in co-cultures initiated with wild-type + G. metallireducens + explanation: Wild-type baseline for the delay, measured in the G. metallireducens-M. barkeri + ethanol coculture that this record describes. + - reference: PMID:37650614 + supports: SUPPORT + evidence_source: IN_VITRO + snippet: In contrast, there was very little lag in co-cultures initiated with the Hcp-deficient + G. metallireducens strain + explanation: Knockout arm of the perturbation - removing the T6SS structural protein Hcp + largely removes the lag, which is the basis for the negative causal edge. + - reference: PMID:37650614 + supports: SUPPORT + evidence_source: IN_VITRO + snippet: The hcp-deficient mutant also established DIET quicker with Methanosarcina barkeri + explanation: Confirms the phenotype specifically for the M. barkeri partner of this community. + - reference: PMID:37650614 + supports: PARTIAL + evidence_source: IN_VITRO + snippet: the mutant also reduced Fe(III) oxide faster than the wild-type strain, a phenotype + not expected from the loss of the T6SS + explanation: Records the pleiotropy confounder - the reason this edge is curated as + T6SS-associated rather than T6SS-caused. environmental_factors: - name: Anaerobic Conditions value: Strict anaerobic diff --git a/kb/communities/Syntrophobacter_Methanobacterium_Syntrophy.yaml b/kb/communities/Syntrophobacter_Methanobacterium_Syntrophy.yaml index 0435f25f..0887a03e 100644 --- a/kb/communities/Syntrophobacter_Methanobacterium_Syntrophy.yaml +++ b/kb/communities/Syntrophobacter_Methanobacterium_Syntrophy.yaml @@ -79,13 +79,24 @@ taxonomy: - SYNTROPHIC_PARTNER abundance_value: H2/formate-utilizing methanogen - syntrophic partner evidence: - - reference: doi:10.1099/00207713-48-4-1383 + - reference: PMID:29611893 supports: SUPPORT evidence_source: IN_VITRO + snippet: We performed a proteome analysis of S. fumaroxidans growing with propionate axenically + with sulfate or fumarate, and in syntrophy with Methanospirillum hungatei, Methanobacterium + formicicum or Desulfovibrio desulfuricans + explanation: Exact-pair evidence that S. fumaroxidans grows syntrophically on propionate with + M. formicicum, the partner of this record. + - reference: doi:10.1099/00207713-48-4-1383 + supports: PARTIAL + evidence_source: IN_VITRO snippet: It oxidized propionate syntrophically in co-culture with the hydrogen- and formate-utilizing Methanospirillum hungateii, and was able to oxidize propionate and other organic compounds in pure culture with sulfate or fumarate as the electron acceptor - explanation: Establishes M. formicicum as the H2/formate-utilizing syntrophic partner + explanation: Establishes S. fumaroxidans as a syntrophic propionate oxidizer that depends on an + H2/formate-utilizing methanogenic partner. PARTIAL - the methanogen named in this passage is + M. hungateii, NOT this record's partner M. formicicum, so it supports the partner ROLE but not + the partner IDENTITY. See discussion kg-syntrophobacter-methanobacterium-partner-attribution. ecological_interactions: - name: Propionate Oxidation and H2/Formate Production description: 'Syntrophobacter fumaroxidans oxidizes propionate to acetate via the methylmalonyl-CoA @@ -186,15 +197,31 @@ ecological_interactions: term: id: GO:0015948 label: methanogenesis + downstream: + - target: Propionate Oxidation and H2/Formate Production + description: H2/formate scavenging by M. formicicum keeps both carriers at low partial + pressure, which is what makes the thermodynamically unfavourable propionate oxidation by + S. fumaroxidans feasible. S. fumaroxidans oxidizes propionate either in syntrophy with an + H2/formate-utilizing methanogen or, axenically, only with an external electron acceptor + (sulfate or fumarate). Reciprocal arm of the syntrophic loop; DIRECTLY IMPLIED by the + axenic-vs-syntrophic contrast rather than demonstrated by a partner-removal or + carrier-inhibition perturbation in this exact pair. evidence: - - reference: doi:10.1099/00207713-48-4-1383 + - reference: PMID:29611893 supports: SUPPORT evidence_source: IN_VITRO + snippet: Syntrophobacter fumaroxidans is a sulfate-reducing bacterium able to grow on propionate + axenically or in syntrophic interaction with methanogens or other sulfate-reducing bacteria + explanation: The axenic-versus-syntrophic contrast underlying the feedback edge - without a + partner, propionate growth requires an external electron acceptor. + - reference: doi:10.1099/00207713-48-4-1383 + supports: PARTIAL + evidence_source: IN_VITRO snippet: It oxidized propionate syntrophically in co-culture with the hydrogen- and formate-utilizing Methanospirillum hungateii, and was able to oxidize propionate and other organic compounds in pure culture with sulfate or fumarate as the electron acceptor - explanation: Demonstrates successful syntrophic coculture with M. formicicum as H2/formate-utilizing - partner + explanation: Demonstrates the syntrophic-coculture dependence on an H2/formate-utilizing methanogen. + PARTIAL - the methanogen named here is M. hungateii, NOT this record's partner M. formicicum. - reference: PMID:29611893 supports: SUPPORT evidence_source: IN_VITRO @@ -319,3 +346,50 @@ growth_media: Methanobacterium formicicum MF T (DSM 1535) were grown with 30 mM of propionate without electron acceptor explanation: Confirms the S. fumaroxidans-M. formicicum coculture and propionate substrate concentration. +discussions: +- discussion_id: kg-syntrophobacter-methanobacterium-partner-attribution + prompt: > + Which curated claims about this consortium rest on exact-pair + (S. fumaroxidans + M. formicicum) evidence, and which were imported from + S. fumaroxidans + M. hungatei experiments? + kind: KNOWLEDGE_GAP + status: OPEN + attaches_to: + - ecological_interactions#Propionate Oxidation and H2/Formate Production + - ecological_interactions#Interspecies Electron Transfer and Methanogenesis + rationale: > + Several evidence items on this record quote doi:10.1099/00207713-48-4-1383 + (Harmsen et al. 1998), whose syntrophic coculture passage names + *Methanospirillum hungateii*, not this record's partner *Methanobacterium + formicicum*. Two of those items previously carried explanations asserting that + the passage established M. formicicum as the partner; they have been corrected + and downgraded to PARTIAL, and exact-pair support has been added from + PMID:29611893 (Sedano-Nunez et al. 2018), which explicitly grew S. fumaroxidans + in syntrophy with M. formicicum. The residual gap: the frequently cited + biochemical formate-transfer and FDH/hydrogenase transcription results for this + system were obtained with M. hungatei, so carrier apportionment ("formate is the + dominant carrier") must NOT be curated as an exact-pair finding. A 2024 + exact-pair perturbation study (Li et al., Water 16:3551, + doi:10.3390/w16243551) reports graded-formate dosing on the + S. fumaroxidans-M. formicicum coculture - low formate (5-10 mM) promoting and + >=30 mM inhibiting syntrophic propionate oxidation, with FDH/hydrogenase + transcript downregulation - which would materially enrich this record's causal + graph. It is not ingested here because the journal is not indexed in PubMed and + has no PMC record, so its snippets cannot be verified by the repo's + reference-cache tooling. + evidence: + - reference: PMID:29611893 + supports: SUPPORT + evidence_source: IN_VITRO + snippet: We performed a proteome analysis of S. fumaroxidans growing with propionate axenically + with sulfate or fumarate, and in syntrophy with Methanospirillum hungatei, Methanobacterium + formicicum or Desulfovibrio desulfuricans + explanation: The one curated reference that studies the exact pair alongside the M. hungatei + pairing - the basis for separating exact-pair from imported claims. + - reference: doi:10.1099/00207713-48-4-1383 + supports: PARTIAL + evidence_source: IN_VITRO + snippet: It oxidized propionate syntrophically in co-culture with the hydrogen- and formate-utilizing + Methanospirillum hungateii + explanation: The misattribution source - this passage names M. hungateii and cannot establish + M. formicicum as the partner. diff --git a/kb/communities/Syntrophobacter_Methanospirillum_Syntrophy.yaml b/kb/communities/Syntrophobacter_Methanospirillum_Syntrophy.yaml index 65efd87a..552b9371 100644 --- a/kb/communities/Syntrophobacter_Methanospirillum_Syntrophy.yaml +++ b/kb/communities/Syntrophobacter_Methanospirillum_Syntrophy.yaml @@ -186,6 +186,14 @@ ecological_interactions: term: id: GO:0015948 label: methanogenesis + downstream: + - target: Propionate Oxidation and H2/Formate Production + description: H2/formate scavenging by M. hungatei keeps the partial pressures of both carriers + low enough for propionate oxidation to remain thermodynamically feasible. S. fumaroxidans + oxidizes propionate either syntrophically with an H2/formate-utilizing partner or, in pure + culture, only when supplied with an external electron acceptor (sulfate or fumarate) — + establishing the partner-dependence that this feedback edge encodes. Reciprocal arm of the + syntrophic loop. evidence: - reference: PMID:9828440 supports: SUPPORT @@ -194,7 +202,8 @@ ecological_interactions: Methanospirillum hungateii, and was able to oxidize propionate and other organic compounds in pure culture with sulfate or fumarate as the electron acceptor explanation: Demonstrates successful syntrophic coculture with M. hungatei as H2/formate-utilizing - partner + partner; the pure-culture contrast (electron acceptor required in the absence of the partner) is + the basis for the feedback edge - reference: PMID:29611893 supports: SUPPORT evidence_source: IN_VITRO diff --git a/kb/communities/Syntrophomonas_Methanospirillum_Syntrophy.yaml b/kb/communities/Syntrophomonas_Methanospirillum_Syntrophy.yaml index 4fd018d3..7c60cfff 100644 --- a/kb/communities/Syntrophomonas_Methanospirillum_Syntrophy.yaml +++ b/kb/communities/Syntrophomonas_Methanospirillum_Syntrophy.yaml @@ -156,7 +156,26 @@ ecological_interactions: term: id: GO:0015948 label: methanogenesis + downstream: + - target: Butyrate Oxidation and H2 Production + description: H2 scavenging by M. hungatei keeps the H2 partial pressure low enough for + butyrate β-oxidation to remain thermodynamically feasible; S. wolfei grows and degrades + butyrate only in association with an H2-using partner, and adding H2 back to the medium + stops both. Reciprocal (feedback) arm of the syntrophic loop. evidence: + - reference: PMID:16345745 + supports: SUPPORT + evidence_source: IN_VITRO + snippet: Growth and degradation of fatty acids occur only in syntrophic association with + H(2)-using bacteria + explanation: Establishes the methanogen-side dependency underlying the feedback edge - S. + wolfei is obligately dependent on an H2-scavenging partner + - reference: PMID:16345745 + supports: SUPPORT + evidence_source: IN_VITRO + snippet: The addition of H(2) to the medium stopped growth and butyrate degradation by S + explanation: Perturbation evidence for the feedback edge - raising H2 (i.e. removing the + effect of H2 scavenging) halts butyrate oxidation - reference: PMID:16345745 supports: SUPPORT evidence_source: IN_VITRO diff --git a/references_cache/PMID_29611893.md b/references_cache/PMID_29611893.md index adf838a0..86fa698b 100644 --- a/references_cache/PMID_29611893.md +++ b/references_cache/PMID_29611893.md @@ -75,4 +75,782 @@ Microbiology and John Wiley & Sons Ltd. DOI: 10.1111/1462-2920.14119 PMCID: PMC5947623 -PMID: 29611893 [Indexed for MEDLINE] \ No newline at end of file +PMID: 29611893 [Indexed for MEDLINE] + +===== OPEN-ACCESS FULL TEXT (Europe PMC PMC5947623) ===== + +379 blackwellopen Environmental Microbiology Environ Microbiol PMC5947623 5947623 5947623 29611893 10.1111/1462-2920.14119 Comparative proteome analysis of propionate degradation by Syntrophobacter fumaroxidans in pure culture and in coculture with methanogens Sedano‐Núñez Vicente T 1 Boeren Sjef 2 Stams Alfons J M 1 3 Plugge Caroline M 1 ✉ 1 Laboratory of Microbiology, Wageningen University & Research, Stippeneng 4, Wageningen, The Netherlands 2 Laboratory of Biochemistry, Wageningen University & Research, Stippeneng 4, Wageningen, The Netherlands 3 Centre of Biological Engineering, University of Minho, Campus de Gualtar, Braga, 4710‐057, Portugal * For correspondence. E‐mail caroline.plugge@wur.nl ; Tel. (+31) 317 483 752. ✉ Corresponding author. 23 4 2018 20 5 1842 1842–1856 17 5 2018 © 2018 The Authors. Environmental Microbiology published by Society for Applied Microbiology and John Wiley & Sons Ltd. This is an open access article under the terms of the http://creativecommons.org/licenses/by-nc-nd/4.0/ License, which permits use and distribution in any medium, provided the original work is properly cited, the use is non‐commercial and no modifications or adaptations are made. Summary +Syntrophobacter fumaroxidans is a sulfate‐reducing bacterium able to grow on propionate axenically or in syntrophic interaction with methanogens or other sulfate‐reducing bacteria. We performed a proteome analysis of S. fumaroxidans growing with propionate axenically with sulfate or fumarate, and in syntrophy with Methanospirillum hungatei, Methanobacterium formicicum or Desulfovibrio desulfuricans. Special attention was put on the role of hydrogen and formate in interspecies electron transfer (IET) and energy conservation. Formate dehydrogenase Fdh1 and hydrogenase Hox were the main confurcating enzymes used for energy conservation. In the periplasm, Fdh2 and hydrogenase Hyn play an important role in reverse electron transport associated with succinate oxidation. Periplasmic Fdh3 and Fdh5 were involved in IET. The sulfate reduction pathway was poorly regulated and many enzymes associated with sulfate reduction (Sat, HppA, AprAB, DsrAB and DsrC) were abundant even at conditions where sulfate was not present. Proteins similar to heterodisulfide reductases (Hdr) were abundant. Hdr/Flox was detected in all conditions while HdrABC/HdrL was exclusively detected when sulfate was available; these complexes most likely confurcate electrons. Our results suggest that S. fumaroxidans mainly used formate for electron release and that different confurcating mechanisms were used in its sulfidogenic metabolism. status released display-pdf yes is-in-collection-domain yes is-olf no is-manuscript no is-preprint no is-journal-matter no is-scanned no is-retracted no Received 2017 Sep 5; Revised 2018 Mar 20; Accepted 2018 Mar 24; Issue date 2018 May. Introduction +Syntrophobacter fumaroxidans is a sulfate‐reducing deltaproteobacterium able to grow on propionate in syntrophy with methanogens (Harmsen et al., 1998 ). It can also grow axenically by fermenting fumarate (Stams et al., 1993 ). To degrade propionate, it requires fumarate or sulfate as electron acceptors, or a H2‐ and formate‐consuming partner in the absence of an electron acceptor. S. fumaroxidans uses the methylmalonyl‐CoA (MMC) pathway to degrade propionate to acetate and CO2 (Plugge et al., 1993 ). Under standard conditions, propionate oxidation to H2, formate and acetate is an endergonic process. Reducing equivalents at the redox levels of reduced ferredoxin (Fdred) and NADH are released in the pyruvate and malate oxidation steps of the pathway respectively. Succinate oxidation via menaquinone is endergonic since the midpoint potential of succinate is more positive (+30 mV) than the menaquinone (−80 mV). Therefore, the reaction requires a transmembrane proton gradient to function (Plugge et al., 2012 ). For this, it has been proposed that S. fumaroxidans uses a periplasmic formate dehydrogenase, cytochrome b:quinone oxidoreductases, the menaquinone loop and a cytoplasmic fumarate reductase to drive energy dependent succinate oxidation (Müller et al., 2010 ). To keep the pathway functioning, the reduced electron mediators need to be re‐oxidized by reducing protons to H2 or CO2 to formate. Consequently, the role of the hydrogen/formate scavenger in the syntrophic association with S. fumaroxidans is to maintain H2 and formate at sufficiently low levels so that propionate degradation becomes energetically feasible (Stams and Dong, 1995 ). The minimal hydrogen partial pressure (pH2) that methanogens can maintain is between 1 and 10 Pa (Thauer et al., 2008 ). This level is not low enough to overcome the most energy‐consuming step in the MMC pathway, the oxidation of succinate to fumarate. To couple this step to proton or CO2 reduction would require a pH2 of 10−10 Pa and a formate concentration below 1 µM (Schink, 1997 ). Therefore, to drive this reaction, the input of metabolic energy is required. An investment of two‐thirds of an ATP via a mechanism known as reverse electron transport (RET) has been suggested by some authors (Van Kuijk et al., 1998a ; Schink and Stams, 2013 ). During RET energy is invested in the form of ATP to generate a proton gradient across the membrane which allows succinate oxidation to proceed (Stams and Plugge, 2009 ). Membrane‐associated proteins, such as ferredoxin:NAD+ oxidoreductases, cytochromes and periplasmic formate dehydrogenases and hydrogenases, have been reported to be involved in RET (Sieber et al., 2012 ; Grein et al., 2013 ). Moreover, novel energy conversion mechanisms have been discovered in anaerobic microorganisms, for instance flavin‐based electron bifurcation and its reversal, electron confurcation (Li et al., 2008 ; Buckel and Thauer, 2013 ; Schink, 2015 ). Genome analyses of S. fumaroxidans revealed membrane associated proteins, such as a fumarate reductase and a Rnf complex, as well as confurcating hydrogenases and formate dehydrogenases possibly involved in energy conservation mechanisms (Müller et al., 2010 ; Pereira et al., 2011 ; Plugge et al., 2012 ; Worm et al., 2014 ). Subsequently, transcriptomics studies with S. fumaroxidans in syntrophic and axenic cultures showed that a periplasmic formate dehydrogenase (Fdh2) and a hydrogenase (Hyn) play an important role to make the endergonic oxidation of succinate possible (Worm et al., 2011a ). Moreover, it was found that confurcating hydrogenases and confurcating formate dehydrogenases (Hyd1, Hox and Fdh1) are important energy conversion enzymes required for propionate degradation (Worm et al., 2011a , 2011b ). In this study, a comparative proteomic analysis of S. fumaroxidans was made. Cells grown with propionate coupled to fumarate or sulfate reduction, or in syntrophic associations with Methanospirillum hungatei or Methanobacterium formicicum were compared. We aim to elucidate the main metabolic differences in lifestyles by identifying the key proteins used by S. fumaroxidans in interspecies electron transfer (IET), reverse electron transport (RET), electron confurcating processes and other energy conservation pathways. In addition to the known syntrophic interactions of S. fumaroxidans with methanogens, our study was extended by including the proteomic profiling of S. fumaroxidans in coculture with a non‐methanogenic partner. Desulfovibrio desulfuricans has been studied before in cocultures with Syntrophobacter wolinii and S. fumaroxidans as a hydrogen‐ or formate‐scavenger in the oxidation of propionate (Boone and Bryant, 1980 ; Dong et al., 1994 ). However, the nature of the symbiotic interactions of such cocultures was not properly defined. S. wolinii and S. fumaroxidans are both able to couple propionate oxidation to sulfate reduction instead of proton reduction (Wallrabenstein et al., 1994 ; Van Kuijk and Stams, 1995 ). D. desulfuricans is a sulfate reducer that utilizes lactate, ethanol, hydrogen and formate in the presence of sulfate, but not acetate, propionate, butyrate or glucose (McInerney et al., 1979 ). Therefore, a syntrophic relationship with S. fumaroxidans, in which hydrogen and formate are produced, would be beneficial for D. desulfuricans. Nonetheless, it is intriguing why Syntrophobacter would engage in a syntrophic association while having sufficient sulfate to grow independently. By comparing the proteomic profile of S. fumaroxidans grown in coculture with D. desulfuricans with the proteomic profiles of the other known syntrophic lifestyles, and the sulfidogenic condition, we expect to be able to define the symbiotic relationship of S. fumaroxidans with D. desulfuricans. Moreover, in a syntrophic coculture with Methanobrevibacter arboriphilus AZ, D. desulfuricans oxidized formate and transferred hydrogen to the methanogenic partner (Dolfing et al., 2008 ). The proteomic analysis of D. desulfuricans growing with hydrogen, formate and in coculture with S. fumaroxidans will reveal further insight into sulfate‐reducing syntrophic cocultures. Results Proteomic overview of S. fumaroxidans and most abundant proteins in all growth conditions The genome of S. fumaroxidans contains 4098 protein coding genes (Plugge et al., 2012 ). Our proteomic analysis accurately identified a total of 813 proteins in the five studied conditions. Of these, 84 were designated as proteins with unknown function. About 514 proteins were detected in all the studied conditions. This core proteome represented slightly more than 60% of all the detected proteins (Supporting Information Fig. S1A). Principal component analysis (PCA) revealed that the protein abundance patterns were reproducible among triplicates of a given growth condition (Supporting Information Fig. S1B). Moreover, it shows that protein patterns of S. fumaroxidans differ depending on the electron acceptor or syntrophic partner used, clearly separating syntrophic methanogenic conditions from the axenic proteomic profiles. Statistical analysis indicated that 509 proteins significantly differed in at least one condition. This means that 304 proteins were constitutively produced in the five analysed conditions. Total intensity‐based absolute quantification (iBAQ) revealed the most abundant proteins produced in the whole analysis (Supporting Information Table S1). Most of these proteins were involved in the methylmalonyl‐CoA pathway, sulfate reduction, electron transfer or energy conservation. Highly abundant proteins under all five conditions included chaperonins (GroEL and GroES), heat shock proteins and ribosomal proteins. Other abundant proteins had annotated functions involved in protection, signalling, transcription and ferrous ion transport. Rubrerythrins and proteins involved in the biosynthesis of cofactors like iron‐molybdenum and molybdopterin were also abundant. Enzymes of the methylmalonyl CoA pathway Previous genomic analyses of S. fumaroxidans predicted several genes coding for proteins involved in the MMC pathway (Müller et al., 2010 ; Plugge et al., 2012 ). Most of these proteins were abundant in our whole‐cell proteome analysis. For those predicted proteins that were not detected, paralogous proteins were found in high levels, which suggest that these proteins have a role in the MMC pathway. For instance, the predicted enzymes for propionate activation (Sfum_3926 to Sfum_3934) and for the conversion of acetyl‐CoA to acetate (Sfum_0388–0389, Sfum_0745–0746, Sfum_1278 and Sfum_3070) were not detected in the present study. Nevertheless, three CoA transferases were detected for the five conditions: CoA‐A (Sfum_0809–0810), CoA‐B (Sfum_0811–0812) and CoA‐S (Sfum_1132–1134) (Fig. 1 ). The amino acid sequences of these proteins indicate a relationship to coenzyme A transferase family I (InterPro IPR004165) and could therefore be involved in propionate activation and/or acetate formation. Figure 1 Relative expression levels of the proteins used in the methylmalonyl‐CoA pathway by Syntrophobacter fumaroxidans. Protein abundance levels are shown after Z‐score normalization. High relative expression is indicated in yellow and low relative expression is indicated in blue. Grey colour means not detected. In the left side the MMC steps are shown levelled to the associated proteins. The rows in the heat map show the detected proteins in five different growth conditions. The columns show from left to right, in triplicates, the electron acceptor used by S. fumaroxidans to couple propionate oxidation: fumarate, sulfate and interspecies compounds transferred to: Desulfovibrio desulfuricans, Methanobacterium formicicum and Methanospirillum hungatei. The asterisk indicates a statistically significant difference in at least one condition. MMC, methylmalonyl‐CoA; Sdh, succinate dehydrogenase; Frd, fumarate reductase; DH, dehydrogenase; CT, carboxyltransferase; ACCT, acetyl‐CoA carboxyltransferase; CoA Trans, coenzyme A transferase. As predicted by previous genome studies (Müller et al., 2010 ; Plugge et al., 2012 ), the main protein complex responsible for the oxidation of succinate to fumarate was the membrane bound succinate dehydrogenase SdhABC (Sfum_1998–2000), which was abundant in all conditions. During axenic growth on propionate with fumarate, S. fumaroxidans converts propionate to succinate. Then, part of the fumarate in this growth condition is oxidized to acetate (Stams et al., 1993 ). This conversion is energy dependent, producing reducing equivalents during malate oxidation and pyruvate decarboxylation, and is only possible by coupling it to the energy yielding reduction of fumarate to succinate. The fumarate reductase FrdABEF (Sfum_4092–4095) complex was detected in higher levels during growth with fumarate. Except for a few subunits, the FrdABEF complex was not detected in cells grown with methanogens as expected since fumarate reduction only occurs when fumarate is provided. However, the complex was consistently detected in cells where sulfate was available, particularly in the coculture with D. desulfuricans. In the genome of S. fumaroxidans two additional gene clusters show similarity to succinate dehydrogenases SdhAB‐1 (Sfum_0172–0174) and SdhAB‐2 (Sfum_2103–2104). SdhAB‐1 was not detected in our study and only the alpha subunit of SdhAB‐2 showed a similar detection profile to FrdABEF. The predicted fumarase in the gene cluster Sfum_2101‐02 was not detected in any condition. Instead, a second fumarase from a non‐clustered gene (Sfum_2336) was abundant in all conditions. The amino acid sequence of this second fumarase corresponds to the previously isolated and characterized class I fumarase from S. fumaroxidans (Van Kuijk et al., 1996 ). Although this protein was abundant in all conditions, lower expression levels were measured in sulfate‐reducing cells. Finally, methylmalonyl‐CoA mutase (Sfum_0458) and succinyl‐CoA synthase (Sfum_1702–1703) were significantly more abundant in syntrophically grown cells, while the pyruvate oxidoreductase (Sfum_2792–2795) showed a lower relative expression during growth with Desulfovibrio. Hydrogenases and formate dehydrogenases involved in electron transfer The genome of S. fumaroxidans indicates the presence of six formate dehydrogenases and eight hydrogenases. Relative abundance levels of the hydrogenases and formate dehydrogenases produced by S. fumaroxidans during propionate degradation under different axenic or cocultured conditions are shown in Fig. 2 . In this figure it can be seen that for most of the detected hydrogenases and formate dehydrogenases, the expression levels measured in syntrophic conditions with methanogens were higher than any of the axenic conditions. From the two predicted periplasmic hydrogenases, Hyn (Sfum_2952‐53) was detected in all conditions albeit more abundant during growth with fumarate and with D. desulfuricans, while Hyd2 (Sfum_0847‐48) was not detected in cells that were grown with D. desulfuricans and only in one triplicate of the sulfate condition. Figure 2 Relative abundance levels of hydrogenases and formate dehydrogenases in Syntrophobacter fumaroxidans during propionate oxidation. Protein abundance levels are shown after Z‐score normalization. The detected proteins are shown for five different growth conditions, in triplicates, according to the electron acceptor used by S. fumaroxidans to oxidize propionate; from left to right: fumarate, sulfate and interspecies compounds transferred to: Desulfovibrio desulfuricans, Methanobacterium formicicum and Methanospirillum hungatei. The colour intensity indicates the degree of protein up‐ or down regulation where high relative expression is indicated in red and low relative expression is indicated in blue; the grey colour represents not detected. Underlined complex names have been predicted to function as confurcating. Locus tags in bold font indicate the catalytic subunit of the complex. The asterisk indicates a statistical significant difference in at least one condition. Proteins for Mvh2 (Sfum_3954‐57) were not found in our analysis, and only the subunits containing the FAD and NAD+‐binding oxidoreductase domains were detected for Mvh1 (Sfum_3535‐37) and Frh (Sfum_2221‐24) so these enzymes were classified as not detected. Of the three cytoplasmic hydrogenases detected, Hox (Sfum_2712‐16) and Fhl‐h (Sfum_1791‐94) were present in all conditions. Lastly, Hyd1 (Sfum_0844) was more abundant in syntrophically grown cells and cells grown with propionate and fumarate, but not when sulfate was present. The three periplasmic formate dehydrogenases (Fdh2, Fdh3 and Fdh5) from S. fumaroxidans were abundant during growth in syntrophy with M. hungatei. However, for syntrophic growth with M. formicicum the detection levels of Fdh5 (Sfum_0035–37) and Fdh3 (Sfum_3509‐11) were significantly lower. Fdh3 was not detected in axenic conditions or in the coculture with D. desulfuricans, and Fdh5 was scarcely detected in such conditions. Cytoplasmic Fdh1 (Sfum_2703‐06) and periplasmic Fdh2 (Sfum_1273‐75) were the most abundant formate dehydrogenases in all conditions. Moreover, significantly higher levels were measured during syntrophic growth. The membrane bound Fhl‐f (Sfum_1795–1806) was abundant in syntrophically grown cells but showed a lower relative expression during axenic growth. Fdh4 (Sfum_0030‐01) had very high relative abundance levels in syntrophic cultures. However, Fdh4 was not detected in the pure culture with fumarate, while only the lowest limits of detection were measured in sulfidogenic growth. The formate transporter (Sfum_2707) was detected in all conditions but more abundant in methanogenic cultures. Redox proteins involved in dissimilatory sulfate reduction A set of proteins required for dissimilatory sulfate reduction have previously been predicted in the genome of S. fumaroxidans (Pereira et al., 2011 ). Sulfate adenylyltransferase (Sat), proton‐translocating pyrophosphatase (HppA), APS reductase (AprAB), dissimilatory sulfite reductase (DsrAB) and DsrC complexes were among the most abundant proteins in all growth conditions. In contrast, neither of the two sulfate transporters (Sfum_0271 and Sfum_0653) predicted in the genome was detected in the analysis. Two periplasmic subunits of the QrcABCD complex (QrcB: Sfum_0610 and QrcC: Sfum_0609) were detected in all conditions and more abundant in syntrophic cultures (Fig. 3 ). The subunit QrcA (Sfum_0611) a membrane‐associated multihaem cytochrome c, was not detected. Sfum_4047 is the only other gene in S. fumaroxidans genome coding for a membrane‐anchored multihaem cytochrome c. The product of this gene was also detected in all conditions and more abundant in the cocultures with M. hungatei and D. desulfuricans. Figure 3 Relative abundance levels of proteins involved in sulfate reduction in Syntrophobacter fumaroxidans. Abundance levels after shown after Z‐score normalization. The columns show in triplicates, the electron acceptor used by S. fumaroxidans to couple propionate oxidation, from left to right: fumarate, sulfate and interspecies compounds transferred to: Desulfovibrio desulfuricans, Methanobacterium formicicum and Methanospirillum hungatei. High relative expression is indicated in red and low relative expression is indicated in blue. Grey colour means not detected. The asterisk indicates a statistical significant difference in at least one condition. The genes coding for the trimeric complex QmoABC (Sfum_1049–1051) are well conserved in all known sulfate‐reducing bacteria (SRB) and are commonly located in a sat‐aprAB‐qmoABC cluster (Pereira et al., 2011 ). Surprisingly, the products of these genes were more abundant in cells grown with fumarate and in syntrophy than in cells grown with sulfate. However, a second QmoABC (Sfum_1285‐87) was detected in the proteome in all conditions. This complex was more abundant in cells grown axenically and in the coculture with M. hungatei. Similarly, the principal subunits of the DsrMKJOP (Sfum_1146–1150) complex were found in all conditions but more abundant in axenic conditions and in the coculture with D. desulfuricans. Heterodisulfide reductases (Hdr) are enzymes present in methanogens and perform the reduction of CoM‐S‐S‐CoB heterodisulfide to CoM‐SH and CoB‐SH (Hedderich et al., 2005 ). Although the substrate of these enzymes CoM‐S‐S‐CoB heterodisulfide has only been found in methanogens, the high number of similar proteins (heterodisulfide reductases‐like) in SRB has been emphasized in several genome analyses (McInerney et al., 2007 ; Strittmatter et al., 2009 ; Pereira et al., 2011 ; Grein et al., 2013 ). Moreover, related enzymes have been purified from other non‐methanogenic archaea (Mander et al., 2004 ). An Hdr was detected in the proteome analysis of S. wolfei (Sieber et al., 2015 ), suggesting that the presence in the genome and production of such an enzyme complex is not dependent of a sulfate‐reducing lifestyle, but rather to microorganisms specialized in low energy metabolism. Two of the three predicted heterodisulfide reductases‐like enzymes in S. fumaroxidans were detected in this study, one associated with a flavin oxidoreductase complex Hdr/Flox (Sfum_1970–1977) and the other with a multicomplex that includes an HdrL, a MvhD and an FeS electron transfer protein: HdrABC/HdrL (Sfum_0819–0824). The Flox section of Hdr/Flox is produced in all conditions. HdrABC/HdrL was abundant when sulfate was present whereas only the subunits containing FAD/NAD‐binding domains were detected in syntrophic cultures. The fifth heterodisulfide reductase‐like found in the genome of S. fumaroxidans is associated with a pyruvate:Fd oxidoreductase, HdrAL/POR (Sfum_0012–0018); this complex was not detected. Other proteins involved in energy conservation The principle of electron bifurcation was originally proposed for a butyryl‐CoA dehydrogenase/electron transferring flavoprotein complex (Bcd‐Etf) in Clostridium kluyveri (Li et al., 2008 ). Since then three more flavin‐containing complexes capable of electron bifurcation from anaerobic bacteria and archaea have been described: [FeFe]‐hydrogenases (Hyd), transhydrogenases (NfnAB) and [NiFe]‐hydrogenase/heterodisulfide reductases (MvhADG–HdrABC) (Schut and Adams, 2009 ; Kaster et al., 2011 ; Huang et al., 2012 ; Buckel and Thauer, 2013 ). Although S. fumaroxidans is not able to grow on butyrate or crotonate, complexes similar to Bcd/Etf have been predicted from the genome. The acyl‐CoA subunit (Sfum_1371) of one of these complexes was abundant in all conditions, while the Etf subunits (Sfum_1372 and Sfum_1373) were detected in lower levels, and the beta subunit was not detected at all in cells grown in cocultures. A second Etf complex from genes Sfum_0106 and Sfum_0107 was abundant in all conditions at similar levels than the acyl‐CoA subunit from gene Sfum_1371. (Supporting Information Fig. S2) Two additional paralogs coding for Acyl‐CoA/Etf complexes were found in the genome (Sfum_3686‐88 and Sfum_3929–3931), but not detected in our proteomic analysis. Finally, NfnAB (Sfum_2150–2151), another electron‐bifurcating iron‐sulfur flavoprotein commonly present in genomic analyses of sulfate reducers was exclusively detected during growth with fumarate. Proteome generalities of Desulfovibrio desulfuricans The complete genome of Desulfovibrio desulfuricans strain G11 has recently become available (Sheik et al., 2017 ). The genome counts with 2892 protein‐coding genes. Our proteome analysis successfully detected 779 proteins among the three growing conditions. The core proteome of D. desulfuricans consists of 317 proteins detected in all studied conditions (Supporting Information Fig. S3A). All these 317 proteins were detected in cells grown in coculture with S. fumaroxidans, while the cells growing with hydrogen or formate yielded more than 750 proteins each. Differences in the proteome composition were explored using PCA (Supporting Information Fig. S3B). The first principal component (PC1; ∼ 70% of total variance) clearly separates growth in coculture from axenic growth in formate or hydrogen. However, PC1 did not establish a difference between growth on hydrogen or on formate. The second principal component (PC2) differentiates the three proteomic profiles, albeit PC2 accounts only for 10% of the variability of the data. Although fewer D. desulfuricans proteins were detected in cells grown in coculture with S. fumaroxidans, proteins required for sulfate reduction such as AprA (G11_01440) were found among the ten most abundant proteins along with a periplasmic formate dehydrogenase (FDH3; G11_11530–11545) and a periplasmic [NiFe]‐hydrogenase (Hyd‐3; G11_06350–06355) (Supporting Information Fig S4). Periplasmic FDH3 and Hyd‐3 were in fact detected in all conditions, as well as cytoplasmic [NiFe]‐Hyd‐1 (G11_01905–01920) and FDH1 (G11_05250–05260) (Supporting Information Fig. S5). The cytoplasmic formate dehydrogenase FDH2 (G11_10090–10100) on the other hand, was detected only in cells grown with formate, while the periplasmic [FeFe]‐hydrogenase Hyd‐4 (G11_09530–09535) and cytoplasmic Ni‐Fe Hyd‐2 and Hyd‐6 (G11_02760‐80 and G11_10370‐75) were found in both axenic conditions but not in cells grown in coculture. Another [NiFe]‐hydrogenase (Hyd‐5; G11_10035‐45) with a cytochrome type‐b domain has been predicted from D. desulfuricans genome, but neither this protein nor the formate transporter (G11_05695) were detected in the proteomic results. Discussion The majority of the most abundant proteins detected in this study were involved in major processes such as propionate degradation, sulfate reduction, electron transfer, and energy conservation. Other abundant proteins, such as heat‐shock proteins, chaperonins, histones and transporters, emphasize the importance of protection, transport and stabilization of diverse macromolecules in the cell. These proteins have previously been reported as highly abundant in several proteomic analyses and identified as common stress‐induced molecules required for normal cell growth (Hemmingsen et al., 1988 ; Lu et al., 2007 ; Mancuso et al., 2012 ; Sieber et al., 2015 ). Energy‐dependent succinate oxidation in MMC For propionate degradation with fumarate, S. fumaroxidans requires a fumarate reductase, whereas to oxidize propionate with sulfate, or in syntrophy, a succinate dehydrogenase is needed. The high levels of the fumarate reductase (FrdABEF) in cells grown with propionate and fumarate reflects the reduction of fumarate in this lifestyle. However, the abundance of this complex in cells growing with sulfate and in coculture with D. desulfuricans can only be explained by a reversible performance to succinate oxidation, since no succinate was accumulated in those conditions. Fumarate reductases and succinate dehydrogenases are functionally and structurally related enzymes (Mattevi et al., 1999 ). The membrane bound SdhABC of S. fumaroxidans has previously been purified, characterized and showed activity in both directions, fumarate reduction and succinate oxidation (Van Kuijk, 1998b ). However, FrdABEF has not been purified and as such could not be tested for a reversible activity. Transcription experiments reported that FrdABEF was up‐regulated (> 2 log ratio) when fumarate was the electron acceptor in contrast with the gene transcription of cells gown in syntrophic cocultures with M. hungatei (Worm, 2010 ). Interestingly in such study FrdABEF was also up‐regulated in cells grown with sulfate as the electron acceptor and down‐regulated in cells cocultured with M. formicicum. Our proteomic study confirms the high expression levels of FrdABEF in propionate plus fumarate cultures. Moreover, FrdABEF was also present in conditions where propionate was oxidized with sulfate and in coculture with D. desulfuricans. These results might suggest a reversible function of the fumarate reductase FrdABEF toward succinate oxidation. Nevertheless, although in in‐vitro analysis the reversible activity of enzymes is possible, in vivo the enzymes are usually dedicated to one physiological function. Besides S. fumaroxidans has a succinate dehydrogenase (SdhABC) for succinate oxidation. A more likely possibility is that fumarate reduction occurred in the sulfidogenic condition. To pull the oxidation of succinate toward the formation of fumarate, hydrogen and formate, these products have to be efficiently removed. To maintain the levels of fumarate low, the fumarase has to convert fumarate efficiently to malate. This process is very important and as such fumarase is one of the most abundant proteins in S. fumaroxidans. However, cells grown with sulfate show the lowest expression levels of this enzyme. It might be that if fumarate is not removed efficiently in sulfate‐grown cells, the bacteria start to produce FrdABEF. Hydrogen and formate in IET and RET During syntrophic growth, S. fumaroxidans needs to transfer electrons via hydrogen and/or formate to a syntrophic partner. It has long been speculated that formate plays a more important role than hydrogen as an electron carrier in the syntrophic associations of this bacterium with methanogens (De Bok et al., 2002a , 2002b ). Although slightly higher levels were measured in the formate transporter during syntrophic growth over the axenic conditions, S. fumaroxidans must rely on other mechanisms to transfer formate. Three formate dehydrogenases (Fdh2, Fdh3 and Fdh5) contain a twin‐arginine translocation (Tat) pathway conserved site, which points to the translocation of these proteins across the cytoplasmic membrane. Fdh3 and Fdh5 were detected only in syntrophically grown cells, while Fdh2 was detected in all conditions, but was more abundant during syntrophic growth. This suggests that periplasmic Fdh3 and Fdh5 are complexes specialized in transferring formate to the syntrophic partner, while Fdh2 is broadly used for energy conservation purposes as part of the reverse electron transport mechanism, possibly coupled to SdhABC or FrdABEF (Fig. 4 ). Figure 4 Schematic representation of energy converting complexes and proteins involved in sulfate reduction in Syntrophobacter fumaroxidans during propionate oxidation. Among the cytoplasmic formate dehydrogenases, Fdh1 is homologous to the bifurcating [FeFe]‐hydrogenase of Thermotoga maritima (Schut and Adams, 2009 ). Furthermore, it contains a conserved site coding for a 51 kDa subunit of a NADH:ubiquinone oxidoreductase which makes this protein a very plausible candidate for a confurcating‐type of formate dehydrogenase. Fdh1 was detected in all conditions and higher levels were detected in syntrophic conditions. Similarly, the membrane associated Fhl‐f was also detected in all conditions and more abundant in syntrophically grown cells. The ubiquitous detection of Fdh1 and Fhl‐f indicates that their role is not restricted to IET, but that these complexes are essential for energy conservation and formate/hydrogen interconversion during propionate degradation. On the other hand, Fdh4 was not detected in cells grown with fumarate, scarcely detected in cells grown with sulfate and highly abundant in methanogenic conditions. This led us to speculate that Fdh4 has an exclusive role in IET. Furthermore, the genes coding for Fdh4 are located upstream in the genome of the periplasmic Fdh5 operon. Considering these observations, we propose that these neighbouring genes coding for cytoplasmic and periplasmic formate dehydrogenases are used mainly for interspecies formate transfer. Thus Fdh3, Fdh4 and Fdh5 seem to form a set of formate dehydrogenases used by S. fumaroxidans to transfer electrons to the syntrophic partner. It is conceivable that these formate dehydrogenases contain a molybdenum catalytic core (Mo‐FDH) in contrast to Fdh1 and Fdh2 whose structure has been characterized and were shown to have only tungsten‐containing active sites (W‐FDH) (De Bok et al., 2003 ). Further biochemical analysis of these formate dehydrogenases will give insight of the role of molybdenum in IET mechanisms in methanogenic environments (Plugge et al., 2009 ; Worm et al., 2011b ). Only five of the eight predicted hydrogenases of S. fumaroxidans were detected in the present analysis. Of the two periplasmic hydrogenases, Hyn was more abundant in cells grown with propionate and fumarate and in coculture with D. desulfuricans. Hyn has been proposed to be involved in reverse electron transport coupled with FrdABEF for fumarate reduction or SdhABC for succinate oxidation (Worm et al., 2011a ). Considering the high levels of Hyn and FrdABEF in the coculture with D. desulfuricans, we suggest that indeed Hyn is involved in RET with FrdABEF, whether reducing fumarate in fumarate conditions or reversibly oxidizing succinate in the coculture with D. desulfuricans (Fig. 4 ). Of the three cytoplasmic hydrogenases detected, Hox and Fhl‐h, which were detected in all conditions, were more abundant in cells grown with the methanogens. Hox is most probably a confurcating hydrogenase involved in energy conservation. The membrane‐bound Fhl‐h on the other hand, together with Fhl‐f might be involved in a cytoplasmic hydrogen‐formate interconversion during syntrophic growth to control electron release. Finally, the genes coding for Hyd1 and Hyd2 are adjacent in the genome, the products of these genes are produced only in the presence of fumarate and during syntrophic growth but not when sulfate was available. This might be due to the exclusive use of other confurcating energy‐conserving complexes in sulfidogenic conditions, for instance HdrABC/HdrL. Although formate formation seems to prevail in the syntrophic lifestyle of S. fumaroxidans, our results indicate that hydrogen, via Hyd1, Hyd2, Hox and Hyn also plays an important role in energy conservation by RET. During growth with fumarate, when IET is not required, these hydrogenases were detected in higher abundance than any of the formate dehydrogenase in such growth condition. Energy conservation mechanisms in the sulfate‐reducing metabolism All the proteins necessary for sulfate reduction in S. fumaroxidans were abundant in this analysis, with the intriguing exception of the sulfate transporters that were not detected. In order to activate sulfate by sulfate adenylyltransferase, sulfate has to be transported into the cell. Therefore, another mechanism for transport of sulfate across the membrane must be used by S. fumaroxidans. Several transporters and unknown proteins were among the most abundant proteins in this study, it is possible that some of them could have played a role in the import of sulfate to the cytoplasm. The abundance of HppA, Sat, Apr and DsrAB in our proteomic analysis in conditions where sulfate reduction was not observed indicates that the sulfate reduction pathway is not strictly regulated in S. fumaroxidans. However, all these enzymes were significantly more abundant in conditions where sulfate was available, indicating sulfidogenic activity in cells grown with sulfate and with D. desulfuricans. Similarly, for complexes such as Qmo‐2, DsrMKJOP and Hdr/Flox it is possible to observe an up‐regulation in axenic conditions and in some cases in coculture with D. desulfuricans, while for Qrc and Qmo‐1 higher levels are observed in syntrophically grown cells. These observations suggest that the use of these complexes in electron transfer is not constrained to a sulfidogenic lifestyle, and that they could for instance transfer electrons to periplasmic formate dehydrogenases for IET or to the FrdABEF for RET. Quinone reductase complexes (QrcABCD) are involved in the reduction of the quinone pool in D. vulgaris Hildenborough. Furthermore, it was shown that QrcABCD is reduced by periplasmic hydrogenases and formate dehydrogenase via the cytochrome c3 (subunit A of the complex) (Venceslau et al., 2010 ). Although in D. vulgaris the described role of QrcABCD is to reduce menaquinone with electrons gained from hydrogen or formate oxidation during sulfate reduction, we speculate that a reverse process is feasible. In D. desulfuricans G20, a mutant lacking the qrcB gene was unable to grow with H2 or formate as electron donor, while it grew similarly as the parent strain with lactate (Li et al., 2009 ). Moreover, this mutation also inhibited syntrophic growth with a methanogen in lactate. The higher levels of the QrcABCD of S. fumaroxidans in cells grown in syntrophy might be explained by its involvement in electron transfer to the periplasmic formate dehydrogenases Fdh3 and Fdh5 (Figs. 2 and 3 ). Direct electron transfer from Qmo to Apr to facilitate the reduction of sulfate to sulfite has been reported in Desulfovibrio desulfuricans (Pires et al., 2003 ; Pereira, 2008 ; Duarte et al., 2016 ). In Syntrophobacter, the higher expression levels of the two Qmo complexes in cells grown with fumarate might be due to the use of this membrane bound complex in transferring electrons to FrdABEF for RET. FrdABEF lacks a transmembrane subunit, therefore it has been speculated that it receives electrons from menaquinone via cytochrome b and cytochrome b:quinone oxidoreductases (Müller et al., 2010 ), however these cytochromes were not detected in our study. DsrMKJOP is another highly conserved membrane complex in SRB (Rabus et al., 2015 ). In many Gram‐positive SRB only the cytoplasmic‐facing DsrMK genes are present, suggesting that this is the minimal functional module (Pereira et al., 2011 ). Although in S. fumaroxidans the complete gene set of DsrMKJOP is present, only the essential subunits (DsrMK), and the periplasmic DsrO were detected in our proteomic study. In the heat map shown in Fig. 3 the expression profile of DsrMKO is similar to that of the Hdr/Flox complex. If Hdr/Flox is used in all conditions to confurcate electrons as will be discussed below, DsrMKO might be involved in electron transfer with this complex. HdrABC/FloxABCD, a novel NADH dehydrogenase/heterodisulfide reductase widespread in anaerobic bacteria has been proposed to be involved in flavin‐based electron bifurcation in D. vulgaris Hildenborough (Ramos et al., 2015 ). The Flox proteins (Sfum_1970–1973) of the Hdr/Flox of S. fumaroxidans were constitutively present in all the conditions. Nevertheless, the Hdr‐like complex in the Hdr/Flox cluster have a composition different to the canonical HdrABC. For instance, HdrBC is replaced by the cysteine‐rich containing HdrD (Sfum_1969), which was not detected in our analysis. Furthermore, two hdrA genes are present (Sfum_1974 and Sfum_1977), but only the product of Sfum_1977 was detected. Hdr/Flox could be another confurcating system used by S. fumaroxidans to oxidize NADH during propionate degradation, possibly involved in recycling NAD+ during the reduction of fumarate. However, the conformational changes mentioned above might imply functional differences that need to be further investigated. For the HdrABC/HdrL complex, the hdrABC genes (Sfum_0819–0821) are next to genes coding for a pyridine nucleotide‐disulphide oxidoreductase comprising an HdrL protein (Sfum_0824). HdrL is a large protein containing HdrA and one or two NADH binding domains (Strittmatter et al., 2009 ; Pereira et al., 2011 ). An MvhD protein is encoded in Sfum_0823, but the catalytic hydrogenase subunit MvhA is not present. The amino acid sequence of Sfum_0822 codes for iron‐sulfur domains (4Fe‐4S) commonly found in the beta subunits of hydrogenases or formate dehydrogenases (InterPro, December 2017). Moreover, a BlastP search of the amino acid sequence resulted in significant alignments with sequences of formate dehydrogenases in other SRB. We can only speculate if this HdrABC/HdrL complex is able to use hydrogen, formate or some other compound, but the high detection levels of the complete multimeric complex imply an important function in the sulfate‐reducing metabolism. HdrABC/HdrL was detected only in conditions where sulfate was present, axenically or in the presence of D. desulfuricans. The soluble complex MvhADG/HdrABC has been shown to perform flavin‐based electron bifurcation in methanogens (Thauer et al., 2008 ; Kaster et al., 2011 ). We speculate that HdrABC/HdrL is preferred when sulfate is available, over the confurcating hydrogenase Hyd1 which in turn was highly abundant in cells grown with fumarate as electron acceptor and in syntrophy, but not detected when sulfate was in the medium (Fig. 2 ). The reason for the preference of HdrABC/HdrL under sulfidogenic conditions is unclear. However, it could be related to the substrates used by this complex. The MvhADG/HdrABC in methanogens uses H2 to reduce ferredoxin and heterodisulfide (Kaster et al., 2011 ). It is possible that the exclusive high levels of HdrABC/HdrL in our sulfidogenic conditions correspond to the need of reduction of the so called “bacterial heterodisulfide” DsrC (Venceslau et al., 2014 ). It has been suggested (Venceslau et al., 2014 ), that the protein DsrC could serve as a redox hub, linking oxidation of several substrates to sulfate reduction. Our results with S. fumaroxidans show DsrC as one of the most abundant proteins present in all conditions and significantly more abundant in syntrophy with M. hungatei. The recent discoveries point to the role of DsrC as an electron carrier interacting with DsrAB, DsrMKJOP, Hdr/Flox and HdrABC/HdrL, but it could also connect other enzyme complexes like the fumarate reductase FrdABEF in our model bacterium S. fumaroxidans, which in turn would also explain the detection of FrdABEF in cells grown with sulfate. Proteomic profiling of Desulfovibrio desulfuricans The low amount of D. desulfuricans proteins detected from cells grown in coculture with S. fumaroxidans can be the result of low biomass in such condition. From microscopic observations we know that the ratio of S. fumaroxidans to D. desulfuricans was 2:1 (data not shown). Although normalization of the data performed with MaxQuant allowed us to compare the detected proteins with the other growth conditions where more proteins were identified, we rather focused in analysing the most abundant proteins detected in the coculture condition. The abundance of the periplasmic Hyd‐3 and periplasmic FDH3 in cells grown with S. fumaroxidans indicates that interspecies electron transfer carried by formate and hydrogen was taking place in the coculture. The abundance of the proteins involved in sulfate reduction confirm that D. desulfuricans was actively reducing sulfate for which it certainly needed electron donors which could only come from S. fumaroxidans in such growth condition. This shows a remarkable metabolic tendency of S. fumaroxidans to engage in syntrophic interactions. Conclusions This study shows the importance of formate as electron carrier in IET and RET during syntrophic and axenic growth of Syntrophobacter fumaroxidans. S. fumaroxidans utilizes a specific set of enzymes (Fdh3, Fdh4 and Fdh5) to transfer electrons to the syntrophic partner. Previous isolation and characterization of Fdh1 and Fdh2 have revealed only tungsten‐containing active sites (W‐FDH). Biochemical analysis of the three above mentioned formate dehydrogenases could provide insight of the role of molybdenum‐dependent formate dehydrogenases in syntrophic growth. Fdh2 and Hyn are the periplasmic enzymes used by S. fumaroxidans to recycle hydrogen and formate during RET. While Fdh2 is mainly coupled to Sdh during succinate oxidation, Hyn seems to be coupled to Frd for fumarate reduction in propionate plus fumarate but also for succinate oxidation in other growth conditions. Although the sulfate‐reducing metabolism is poorly regulated, the abundance of membrane‐bound complexes like Qrc, Qmo and DsrMKJOP, consistently found in all conditions, as well as the absence of cytochromes in the present study (only two cytochromes detected from eight predicted in the genome), indicates that those membrane‐bound complexes might play a role in the transfer of electrons between cytoplasmic enzymes and the periplasmic formate dehydrogenases and hydrogen dehydrogenases. HdrABC/HdrL is the most abundant putatively confurcating system in sulfidogenic conditions, possibly because of its probable connection to DsrC, an electron hub in sulfidogenic metabolism. The proteomic profiles of both bacteria in the coculture of S. fumaroxidans with D. desulfuricans give insight in the metabolic flexibility of S. fumaroxidans. Results showed a proteomic profile of S. fumaroxidans in which sulfate reduction took place, while energy conservation and IET mechanisms were also used similarly as in the syntrophic associations with methanogens. The proteomic analysis of the partner D. desulfuricans confirmed IET via formate and hydrogen carried on by S. fumaroxidans in a sulfate rich environment. Materials and methods Organisms and growth conditions +Syntrophobacter fumaroxidans was grown in pure culture and in cocultures. Syntrophobacter fumaroxidans MPOBT (DSM 10017) was cultivated under anoxic conditions in basal medium as described previously (Stams et al., 1993 ). The medium for the pure cultures was supplemented with 20 mM propionate and 60 mM fumarate. Sulfidogenic cultures were grown on 20 mM propionate and 20 mM sulfate. Cocultures of S. fumaroxidans with Methanospirillum hungatei strain JF1T (DSM 864) or Methanobacterium formicicum MFT (DSM 1535) were grown with 30 mM of propionate without electron acceptor. A coculture of S. fumaroxidans with Desulfovibrio desulfuricans strain G11 (DSM 7057; Sheik et al., 2017 ) was grown with 20 mM propionate and 20 mM sulfate. Axenic cultures of D. desulfuricans were grown with 20 mM sulfate and 40 mM formate or hydrogen (1.7 atm H2/CO2 80:20 vol/vol). All organisms were batch cultured in triplicate at 37°C in 1 l flasks with 550 ml medium under anaerobic conditions provided by a pressurised (172 kPa; 1.7 atm) gas phase of N2/CO2 (80:20, vol/vol). Growth was monitored by measuring substrate consumption and product formation (propionate, sulfate, methane, acetate, succinate, malate and/or sulfide). Cells were harvested during mid‐exponential growth phase. The cultures for the experiment were inoculated with cells from cultures that were adapted to these conditions by transferring them at least five times in media with the respective substrates before the start of the experiment. Harvesting cells and Percoll gradient centrifugation Cells were aerobically harvested by centrifugation at 16,000g for 16 min at 4°C. The pellet was washed twice with TE buffer (10 mM Tris‐HCl, pH 7.5; 1 mM EDTA). Only cells from the syntrophic coculture of S. fumaroxidans and M. hungatei were separated by Percoll gradient centrifugation (Percoll®, Sigma‐Aldrich, Missouri) as described elsewhere (De Bok et al., 2002b ). The separated layers, containing Syntrophobacter cells in the upper layer and Methanospirillum cells in the lower layer, were collected and subjected to Percoll gradient separation a second time. Cells were then washed twice with 10 mM sodium phosphate buffer (pH 7.5). Protein extraction and SDS‐PAGE Cells were resuspended in lysis buffer (100 mM Tris‐HCl, pH 7.5; 4% w/v SDS; 50 mM dithiothreitol and SIGMAFAST™ Protease Inhibitor Cocktail Tablet—Sigma‐Aldrich, Missouri), and passed three times through a French press (French® Type Pressure Cell Disrupter, Stansted Fluid Power, Harlow, UK) at 2 MPa (40 K cell). Cell debris and undisrupted cells were removed by centrifugation at 18,000g for 10 min at 4°C. The supernatant was collected in Eppendorf™ LoBind Protein Microcentrifuge Tubes and stored at −80°C. Still in the lysis buffer, proteins were denatured by heating at 95°C for 5 min. Samples were loaded on a 10% polyacrylamide separation gel (Precise™ Tris‐HEPES Gels, Thermo Scientific, Rockford) using the Mini‐PROTEAN Tetra Cell (Bio‐Rad Laboratories B.V, Veenendaal, The Netherlands). The electrophoresis procedure was according to the precast gels manufacturer's instructions. Gels were stained using Coomassie Brilliant Blue (CBB) R‐250. Protein concentration was normalized among triplicates and samples in a qualitative way by analysing the gel pictures taken with G:BOX Chemi XT4 (Syngene, Cambridge, UK) and using the software GeneSys version 1.5.5.0 (GeneTools version 4.03.01). In‐gel trypsin digestion In‐gel digestion of proteins and purification of peptides was done following a modified version of a previously described protocol (Rupakula et al., 2013 ). Disulfide bridges in proteins were reduced by covering the gels with reducing solution (10 mM dithiothreitol, pH 7.6, in 50 mM NH4HCO3), and the gels were incubated at 60°C for 1 h. Alkylation was performed in darkness and shaking (100 rpm) for 1 h by adding 25 ml of iodoacetamide solution (10 mM iodoacetamide in 100 mM Tris‐HCl, pH 8.0). Gels were thoroughly rinsed with demineralized water in between steps. Each gel lane was cut into three slices, and the slices were cut into approximately 1 mm3 cubes and transferred to a separate 0.5 ml protein LoBind tube (Eppendorf, Hamburg, Germany). Enzymatic digestion was done with trypsin sequencing grade (Roche, Mannheim, Germany). About 100 µl of trypsin solution (5 ng µl−1 trypsin in 50 mM NH4HCO3) were added to each tube and incubated 2 h at 45°C with gentle shaking. To stop trypsin digestion, trifluoroacetic acid (10%) was added to the supernatant to lower the pH below 5. The digested protein mixture was purified and concentrated using an in‐house made SPE pipette tip (Lu et al., 2011 ).To recover hydrophobic peptides, 50 µl acetonitrile (vol/vol in 0.1% formic acid) was passed through the column. Finally, the volume was reduced to 20 µl using a SpeedVac concentrator and then adjusted to 50 µl with 0.1% formic acid. Samples were analysed using nLC–MS/MS with a Proxeon EASY nLC and a LTQ‐Orbitrap XL mass spectrometer as previously described (Lu et al., 2011 ). LC–MS data analysis The obtained MS/MS spectra were processed with MaxQuant v. 1.5.2.8. Database with the protein sequences of S. fumaroxidans was downloaded from UniProt ( http://www.uniprot.org ). The protein database of Desulfovibrio desulfuricans strain G11 was downloaded from GenBank accession number CP023415 . An additional dataset with protein sequences of common contaminants (trypsin, human keratins and bovine serum albumin) was included. False discovery rates (FDR) of < 1% were set at peptide and protein levels. Modifications for acetylation (Protein N‐term), deamidation (N, Q) and oxidation (M) were allowed to be used for protein identification and quantification. All other quantification settings were kept default. Filtering and further bioinformatics and statistical analysis were performed with Perseus v.1.5.3.0. Proteins included in our analysis contain at least two identified peptides of which at least one is unique and at least one unmodified. Reversed hits and contaminants were filtered out. Protein groups were filtered to require three valid values in at least one experimental group. Label‐free quantification (LFQ) intensities (values normalized with respect to the total amount of protein and all of its identified peptides) were used to analyse the abundance of proteins in the fractions and further statistical comparisons among conditions. LFQ intensities were transformed to logarithmic values base 10. Missing values were imputed with random numbers from a normal distribution, the mean and standard deviation of which were chosen to best simulate low abundance values close to noise level (Width: 0.3 and downshift 1.8 times). A multiple‐sample test (ANOVA) with permutation‐based FDR statistics (250 permutations, FDR = 0.01 and S0 = 1) was applied to filter significant proteins. PCA were performed with default settings and without category enrichment in components. Z‐score normalization in which the mean of each row (where each row is a protein in triplicate and in different conditions) is subtracted from each value and the result divided by the standard deviation of the row was applied before clustering. Hierarchical clustering of rows, using Euclidean distances, produced a heat map representation of the clustered data matrix. Row clusters were automatically defined (100) and exported to a new matrix. Imputed values were then replaced back to missing values and previously defined clusters were displayed in a new heat map. For D. desulfuricans the Z‐score and hierarchical clustering was done for columns instead of rows in order to compare the most abundant proteins detected in each condition. Supporting information Additional Supporting Information may be found in the online version of this article at the publisher's web‐site: +Fig. S1. A. Venn diagram of the 813 proteins detected in Syntrophobacter fumaroxidans growth on propionate with five different (biological or chemical) electron acceptors. B. Principal Component Analysis performed for S. fumaroxidans protein profiles obtained from each triplicate grown under five different conditions. Symbols: Orange diamonds, sulfate reducing; Red crosses, growth with fumarate; Grey squares, in coculture with Desulfovibrio desulfuricans in a sulfate rich environment; Green triangles, in syntrophy with Methanospirillum hungatei; Blue circles, in syntrophy with Methanobacterium formicicum. +Fig. S2. Normalized expression matrix of energy conservation mechanisms predicted for Syntrophobacter fumaroxidans. Proteins are shown for five different growth conditions, in triplicates; from left to right: fumarate, sulfate and interspecies compounds transferred to: Desulfovibrio desulfuricans, Methanobacterium formicicum and Methanospirillum hungatei. The colour scale illustrates the relative detection level of each protein across the 5 samples; blue (log ratio −2.5) and yellow (log ratio 2.5) indicate lower and higher levels compared with the average level value (in black) respectively. Not detected proteins in a specific condition appear in grey. The asterisk indicates a statistical significant difference in at least one condition. +Fig. S3. A. Venn diagram of the 779 proteins detected in Desulfovibrio desulfuricans growing in sulfate rich medium in coculture with Syntrophobacter fumaroxidans or axenically on H2/CO2 or formate. B. PCA performed for D. desulfuricans protein profiles. Symbols: red diamonds, hydrogenotrophic conditions; black squares, growth with formate and filled grey squares correspond to the cocultured partnership of D. desulfuricans with S. fumaroxidans. +Fig. S4. Heat map of hierarchical clustered proteins produced by Desulfovibrio desulfuricans. The proteins are shown in a clustered matrix after column Z‐score normalization and automatic hierarchical columns clustering. Three growth conditions, in triplicates, are shown according to the electron donor used; from left to right: formate, hydrogen and compounds transferred from Syntrophobacter fumaroxidans. The colour scale represents the relative detection level of each protein across the samples; blue log ratio −3, yellow log ratio 3, red log ratio 4 and green log ratio 5 indicate lower and higher levels compared with the average level value 0 (in black) respectively. The colour intensity indicates the degree of protein up‐ or down regulation; the grey colour represents not detected. +Fig. S5. Normalized expression matrix of hydrogenases and formate dehydrogenases of Desulfovibrio desulfuricans. The rows in the heat map show proteins levels after row Z‐score standardization in three different growth conditions. The columns show from left to right, in triplicates, the electron donor used by D. desulfuricans: formate, hydrogen and interspecies compounds transferred from Syntrophobacter fumaroxidans. The colour scale indicates the degree of protein down‐ or up regulation ranging from blue (−2.2 log ratio), to yellow (2.2 log ratio). The colour intensities indicate lower and higher levels compared with the average level 0 value (in black); the grey colour represents not detected. Subunits, twin‐arginine translocation (TAT) pathway signal and selenocysteine insertion (Sec) sequences are indicated after the locus tag. +Fig. S6. Heat map of hierarchical clustered proteins produced by Syntrophobacter fumaroxidans for propionate degradation. The proteins are shown in a clustered matrix after automatic hierarchical cluster of rows from row Z‐score normalization values. Proteins appear from left to right, in triplicates, according to the growth conditions defined by the electron acceptor used by S. fumaroxidans to oxidize propionate: fumarate, sulfate and interspecies compounds transferred to: Desulfovibrio desulfuricans, Methanobacterium formicicum and Methanospirillum hungatei. The colour scale illustrates the relative detection level of each protein across the samples; blue (log ratio −2.5), yellow (log ratio 2.5) and red (log ratio 3) indicate lower and higher levels compared with the average level value 0 (in black). The colour intensity indicates the degree of protein up‐ or down regulation; the grey colour represents not detected. Click here for additional data file. +Table S1. iBAQ values of proteins detected in axenic and cocultured conditions in Syntrophobacter fumaroxidans and Desulfovibrio desulfuricans. Click here for additional data file. Supporting Information Dataset 1 Click here for additional data file. 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Environ Microbiol +13: 1228–1235. + +Worm, P. +, + +Koehorst, J.J. +, + +Visser, M. +, + +Sedano‐Nunez, V.T. +, + +Schaap, P.J. +, + +Plugge, C.M. +, et al (2014) A genomic view on syntrophic versus non‐syntrophic lifestyle in anaerobic fatty acid degrading communities. Biochim Biophys Acta +1837: 2004–2016. + Associated Data Supplementary Materials Additional Supporting Information may be found in the online version of this article at the publisher's web‐site: +Fig. S1. A. Venn diagram of the 813 proteins detected in Syntrophobacter fumaroxidans growth on propionate with five different (biological or chemical) electron acceptors. B. Principal Component Analysis performed for S. fumaroxidans protein profiles obtained from each triplicate grown under five different conditions. Symbols: Orange diamonds, sulfate reducing; Red crosses, growth with fumarate; Grey squares, in coculture with Desulfovibrio desulfuricans in a sulfate rich environment; Green triangles, in syntrophy with Methanospirillum hungatei; Blue circles, in syntrophy with Methanobacterium formicicum. +Fig. S2. Normalized expression matrix of energy conservation mechanisms predicted for Syntrophobacter fumaroxidans. Proteins are shown for five different growth conditions, in triplicates; from left to right: fumarate, sulfate and interspecies compounds transferred to: Desulfovibrio desulfuricans, Methanobacterium formicicum and Methanospirillum hungatei. The colour scale illustrates the relative detection level of each protein across the 5 samples; blue (log ratio −2.5) and yellow (log ratio 2.5) indicate lower and higher levels compared with the average level value (in black) respectively. Not detected proteins in a specific condition appear in grey. The asterisk indicates a statistical significant difference in at least one condition. +Fig. S3. A. Venn diagram of the 779 proteins detected in Desulfovibrio desulfuricans growing in sulfate rich medium in coculture with Syntrophobacter fumaroxidans or axenically on H2/CO2 or formate. B. PCA performed for D. desulfuricans protein profiles. Symbols: red diamonds, hydrogenotrophic conditions; black squares, growth with formate and filled grey squares correspond to the cocultured partnership of D. desulfuricans with S. fumaroxidans. +Fig. S4. Heat map of hierarchical clustered proteins produced by Desulfovibrio desulfuricans. The proteins are shown in a clustered matrix after column Z‐score normalization and automatic hierarchical columns clustering. Three growth conditions, in triplicates, are shown according to the electron donor used; from left to right: formate, hydrogen and compounds transferred from Syntrophobacter fumaroxidans. The colour scale represents the relative detection level of each protein across the samples; blue log ratio −3, yellow log ratio 3, red log ratio 4 and green log ratio 5 indicate lower and higher levels compared with the average level value 0 (in black) respectively. The colour intensity indicates the degree of protein up‐ or down regulation; the grey colour represents not detected. +Fig. S5. Normalized expression matrix of hydrogenases and formate dehydrogenases of Desulfovibrio desulfuricans. The rows in the heat map show proteins levels after row Z‐score standardization in three different growth conditions. The columns show from left to right, in triplicates, the electron donor used by D. desulfuricans: formate, hydrogen and interspecies compounds transferred from Syntrophobacter fumaroxidans. The colour scale indicates the degree of protein down‐ or up regulation ranging from blue (−2.2 log ratio), to yellow (2.2 log ratio). The colour intensities indicate lower and higher levels compared with the average level 0 value (in black); the grey colour represents not detected. Subunits, twin‐arginine translocation (TAT) pathway signal and selenocysteine insertion (Sec) sequences are indicated after the locus tag. +Fig. S6. Heat map of hierarchical clustered proteins produced by Syntrophobacter fumaroxidans for propionate degradation. The proteins are shown in a clustered matrix after automatic hierarchical cluster of rows from row Z‐score normalization values. Proteins appear from left to right, in triplicates, according to the growth conditions defined by the electron acceptor used by S. fumaroxidans to oxidize propionate: fumarate, sulfate and interspecies compounds transferred to: Desulfovibrio desulfuricans, Methanobacterium formicicum and Methanospirillum hungatei. The colour scale illustrates the relative detection level of each protein across the samples; blue (log ratio −2.5), yellow (log ratio 2.5) and red (log ratio 3) indicate lower and higher levels compared with the average level value 0 (in black). The colour intensity indicates the degree of protein up‐ or down regulation; the grey colour represents not detected. Click here for additional data file. +Table S1. iBAQ values of proteins detected in axenic and cocultured conditions in Syntrophobacter fumaroxidans and Desulfovibrio desulfuricans. Click here for additional data file. Supporting Information Dataset 1 Click here for additional data file. Supporting Information Dataset 2 Click here for additional data file. diff --git a/references_cache/PMID_34939136.txt b/references_cache/PMID_34939136.txt new file mode 100644 index 00000000..66d82c7b --- /dev/null +++ b/references_cache/PMID_34939136.txt @@ -0,0 +1,39 @@ +1. Appl Microbiol Biotechnol. 2022 Jan;106(2):865-876. doi: +10.1007/s00253-021-11736-7. Epub 2021 Dec 23. + +Mechanisms underlying Clostridium pasteurianum's metabolic shift when grown with +Geobacter sulfurreducens. + +Berthomieu R(1), Pérez-Bernal MF(1), Santa-Catalina G(1), Desmond-Le Quéméner +E(1), Bernet N(1), Trably E(2). + +Author information: +(1)INRAE, Univ Montpellier, LBE, Narbonne, France. +(2)INRAE, Univ Montpellier, LBE, Narbonne, France. eric.trably@inrae.fr. + +Recently, a study showed that glycerol fermentation by Clostridium pasteurianum +could be metabolically redirected when the electroactive bacterium Geobacter +sulfurreducens was added in the culture. It was assumed that this metabolic +shift of the fermentative species resulted from an interspecies electron +transfer. The aim of this study was to find out the mechanisms used for this +interaction and how they affect the metabolism of C. pasteurianum. To get +insights into the mechanisms involved, several coculture setups and RNA +sequencing with differential expression analysis were performed. As a result, a +putative interaction model was proposed: G. sulfurreducens produces cobamide +molecules that possibly modify C. pasteurianum metabolic pathway at the key +enzyme glycerol dehydratase, and affect its vanadium nitrogenase expression. In +addition, the results suggested that G. sulfurreducens' electrons could enter C. +pasteurianum through its transmembrane flavin-bound polyferredoxin and cellular +cytochrome b5-rubredoxin interplay, putatively reinforcing the metabolic shift. +Unravelling the mechanisms behind the interaction between fermentative and +electroactive bacteria helps to better understand the role of bacterial +interactions in fermentation setups. KEY POINTS: • C. pasteurianum-G. +sulfurreducens interaction inducing a metabolic shift is mediated • C. +pasteurianum's metabolic shift in coculture might be induced by cobamides • +Electrons possibly enter C. pasteurianum through a multiflavin polyferredoxin. + +© 2021. The Author(s), under exclusive licence to Springer-Verlag GmbH Germany, +part of Springer Nature. + +DOI: 10.1007/s00253-021-11736-7 +PMID: 34939136 [Indexed for MEDLINE] \ No newline at end of file diff --git a/references_cache/PMID_37650614.txt b/references_cache/PMID_37650614.txt new file mode 100644 index 00000000..1fd4e526 --- /dev/null +++ b/references_cache/PMID_37650614.txt @@ -0,0 +1,2079 @@ +1. Microbiol Spectr. 2023 Oct 17;11(5):e0094123. doi: 10.1128/spectrum.00941-23. +Epub 2023 Aug 31. + +Detrimental impact of the Geobacter metallireducens type VI secretion system on +direct interspecies electron transfer. + +Smith JA(#)(1)(2), Holmes DE(#)(1)(3), Woodard TL(1), Li Y(1)(4), Liu X(1)(5), +Wang L-Y(1), Meier D(1), Schwarz IA(2), Lovley DR(1). + +Author information: +(1)Department of Microbiology, University of Massachusetts Amherst, Morrill IV N +Science Center , Amherst, Massachusetts, USA. +(2)Department of Biomolecular Sciences, Central Connecticut State University , +New Britain, Connecticut, USA. +(3)Department of Physical and Biological Sciences, Western New England +University , Springfield, Massachusetts, USA. +(4)School of Ocean Science and Technology, Dalian University of Technology , +Panjin, Liaoning, China. +(5)College of Environmental Science and Engineering, Beijing Forestry University +, Beijing, China. +(#)Contributed equally + +Direct interspecies electron transfer is an alternative to the much more +intensively studied process of interspecies H2 transfer as a mechanism for +microbes to share electrons during the cooperative metabolism of energy sources. +DIET is an important process in anaerobic soils and sediments generating +methane, a significant greenhouse gas. Facilitating DIET can accelerate and +stabilize the conversion of organic wastes to methane biofuel in anaerobic +digesters. Therefore, a better understanding of the factors controlling how fast +DIET partnerships are established is expected to lead to new strategies for +promoting this bioenergy process. The finding that when co-cultured with G. +sulfurreducens, G. metallireducens initially expressed a type VI secretion +system, a behavior not conducive to interspecies cooperation, illustrates the +complexity of establishing syntrophic relationships. + +DOI: 10.1128/spectrum.00941-23 +PMCID: PMC10580878 +PMID: 37650614 + +Conflict of interest statement: The authors declare no conflict of interest. + +===== OPEN-ACCESS FULL TEXT (Europe PMC PMC10580878) ===== + +pmc Microbiol Spectr Microbiol Spectr 3931 microbiolspectr Spectrum Microbiology Spectrum 2165-0497 American Society for Microbiology (ASM) PMC10580878 PMC10580878.1 10580878 10580878 37650614 10.1128/spectrum.00941-23 00941-23 spectrum.00941-23 1 Research Article environmental-microbiology Environmental Microbiology Detrimental impact of the Geobacter metallireducens type VI secretion system on direct interspecies electron transfer https://orcid.org/0000-0003-4248-6566 Smith Jessica A. +1 + +2 + Conceptualization Data curation Formal analysis Investigation Methodology Supervision Writing – original draft Writing – review and editing jsmith@ccsu.edu https://orcid.org/0000-0002-1267-1771 Holmes Dawn E. +1 + +3 + Conceptualization Data curation Formal analysis Investigation Methodology Writing – original draft Writing – review and editing Woodard Trevor L. +1 + Data curation Investigation Supervision Li Yang +1 + +4 + Data curation Formal analysis Investigation Methodology Liu Xinying +1 + +5 + Investigation Methodology Wang Li-Ying +1 + Data curation Formal analysis Investigation Methodology Meier David +1 + Data curation Formal analysis Investigation Schwarz Ingrid A. +2 + Data curation Investigation https://orcid.org/0000-0001-7158-3555 Lovley Derek R. +1 + Conceptualization Supervision Writing – original draft Writing – review and editing + 1 +Department of Microbiology, University of Massachusetts Amherst, Morrill IV N Science Center, Amherst , Massachusetts , USA + + 2 +Department of Biomolecular Sciences, Central Connecticut State University, New Britain , Connecticut , USA + + 3 +Department of Physical and Biological Sciences, Western New England University, Springfield , Massachusetts , USA + + 4 +School of Ocean Science and Technology, Dalian University of Technology, Panjin , Liaoning , China + + 5 +College of Environmental Science and Engineering, Beijing Forestry University, Beijing , China + Editor Afriat-Jurnou Livnat +Migal-Galilee Research Institute, Kiryat Shmona , Israel + Address correspondence to Jessica A. Smith, jsmith@ccsu.edu + Jessica A. Smith and Dawn E. Holmes contributed equally to this article. Author order was determined by a coin toss. The authors declare no conflict of interest. Sep-Oct 2023 31 8 2023 11 5447453 e00941-23 + 02 3 2023 + 02 7 2023 + 31 08 2023 18 10 2023 25 10 2025 Copyright © 2023 Smith et al. 2023 Smith et al. https://creativecommons.org/licenses/by/4.0/ This is an open-access article distributed under the terms of the Creative Commons Attribution 4.0 International license . ABSTRACT Direct interspecies electron transfer (DIET) is important in anaerobic communities of environmental and practical significance. Other than the need for close physical contact for electrical connections, the interactions of DIET partners are poorly understood. Type VI secretion systems (T6SSs) typically kill competitive microbes. Surprisingly, Geobacter metallireducens highly expressed T6SS genes when DIET-based co-cultures were initiated with Geobacter sulfurreducens. T6SS gene expression was lower when the electron shuttle anthraquinone-2,6-disulfonate was added to alleviate the need for interspecies contact. Disruption of hcp, the G. metallireducens gene for the main T6SS needle-tube protein subunit, and the most highly upregulated gene in DIET-grown cells eliminated the long lag periods required for the initiation of DIET. The mutation did not aid DIET in the presence of granular-activated carbon (GAC), consistent with the fact that DIET partners do not make physical contact when electrically connected through conductive materials. The hcp-deficient mutant also established DIET quicker with Methanosarcina barkeri. However, the mutant also reduced Fe(III) oxide faster than the wild-type strain, a phenotype not expected from the loss of the T6SS. Quantitative PCR revealed greater gene transcript abundance for key components of extracellular electron transfer in the hcp-deficient mutant versus the wild-type strain, potentially accounting for the faster Fe(III) oxide reduction and impact on DIET. The results highlight that interspecies interactions beyond electrical connections may influence DIET effectiveness. The unexpected increase in the expression of genes for extracellular electron transport components when hcp was deleted emphasizes the complexities in evaluating the electromicrobiology of highly adaptable Geobacter species. IMPORTANCE Direct interspecies electron transfer is an alternative to the much more intensively studied process of interspecies H2 transfer as a mechanism for microbes to share electrons during the cooperative metabolism of energy sources. DIET is an important process in anaerobic soils and sediments generating methane, a significant greenhouse gas. Facilitating DIET can accelerate and stabilize the conversion of organic wastes to methane biofuel in anaerobic digesters. Therefore, a better understanding of the factors controlling how fast DIET partnerships are established is expected to lead to new strategies for promoting this bioenergy process. The finding that when co-cultured with G. sulfurreducens, G. metallireducens initially expressed a type VI secretion system, a behavior not conducive to interspecies cooperation, illustrates the complexity of establishing syntrophic relationships. KEYWORDS Geobacter type VI secretion system direct interspecies electron transfer extracellular electron transfer Methanosarcina pmc-status-qastatus 0 pmc-status-live yes pmc-status-embargo no pmc-status-released yes pmc-prop-open-access yes pmc-prop-olf no pmc-prop-manuscript no pmc-prop-legally-suppressed no pmc-prop-has-pdf yes pmc-prop-has-supplement yes pmc-prop-pdf-only no pmc-prop-suppress-copyright no pmc-prop-is-real-version no pmc-prop-is-scanned-article no pmc-prop-preprint no pmc-prop-in-epmc yes pmc-license-ref CC BY cover-date September/October 2023 INTRODUCTION A better understanding of the physiological characteristics of microbes that participate in direct interspecies electron transfer (DIET) is required in order to determine how both natural and engineered anoxic environments function ( 1 + – + 3 ). For example, DIET appears to be the primary route for electron exchange between electron-donating bacteria and electron-accepting partners in some types of anaerobic digesters ( 4 , 5 ). In digesters in which interspecies H2 transfer predominates, modifying operating conditions to enhance DIET can accelerate and stabilize the conversion of organic wastes to methane, a needed improvement to this important bioenergy strategy ( 3 , 6 ). Molecular studies have demonstrated that DIET may be a major process in terrestrial methanogenic environments that are significant sources of atmospheric methane ( 7 ), a conclusion that is further supported by the reinterpretation of data on H2 fluxes in these environments ( 8 ). Most of the initial research following the discovery of DIET ( 9 ) focused on identifying which microbes have the potential to participate in DIET and the organic substrates that can support DIET ( 5 , 8 , 10 + – + 21 ). Study of the expression of genes and proteins that enhance electron exchange between species has also been emphasized ( 5 , 9 , 10 , 13 , 22 + – + 25 ). However, other adaptations that promote the switch from a free-living existence to living in close physical association, as is necessary to establish electrical connections for DIET, seem likely. The expression of type VI secretion systems (T6SSs) is expected to be antithetical to interspecies cooperation. Approximately 25% of Gram-negative bacteria have T6SSs that form contractile nanomachines that inject toxins directly into other microbes to eliminate their competition ( 26 + – + 36 ). T6SSs are important in such polymicrobial environments as the human colon ( 37 + – + 41 ), cow rumen ( 42 , 43 ), the plant rhizosphere ( 44 , 45 ), the light organ of the bobtail squid ( 46 ), and soil ( 47 + – + 49 ). In some instances, T6SSs can also be involved in such non-antagonistic behaviors as the modulation of quorum sensing and stress response ( 50 ), self-recognition ( 51 + – + 53 ), and the acquisition of various metals such as zinc, copper, manganese, or iron ( 48 + – + 51 , 54 + – + 57 ). Molecular analyses have demonstrated that Geobacter species are important electron-donating partners for DIET in natural environments, such as subsurface terrestrial soils ( 7 ) as well as in some anaerobic digesters ( 4 , 5 ). The often-observed enrichment of Geobacter when methane production is stimulated with the addition of conductive materials provides further circumstantial evidence for the role of Geobacter in DIET ( 3 , 6 , 58 ). The availability of pure cultures of genetically tractable Geobacter species that can participate in DIET in defined co-cultures has enabled elucidation of important electrical contacts for DIET, such as multi-heme c-type cytochromes and electrically conductive pili ( 5 , 9 , 10 , 23 , 25 , 59 ) as well as strategies for enhancing DIET with electrically conductive minerals and carbon materials ( 60 + – + 64 ). However, Geobacter species are also often found to be free-living in anaerobic soils and sediments, typically transferring electrons to extracellular electron acceptors such as Fe(III) oxides and humic substances ( 65 ). T6SS genes are present in some but not all Geobacter genomes ( 66 , 67 ) (Table S1; Fig. S1). It might be expected that T6SSs could be beneficial to free-living Geobacter species competing against other microbes for resources but not for developing syntrophic cooperation. Here, we report that Geobacter metallireducens highly expresses genes coding for its T6SS in the initial stages of establishing a DIET-based co-culture with Geobacter sulfurreducens, a factor possibly lengthening the adaptation period required for DIET-based growth of the co-culture and accounting for the ability of conductive materials to accelerate DIET. MATERIALS AND METHODS Laboratory strains and culture conditions +Geobacter cultures were obtained from our laboratory culture collection and routinely cultured under strict anaerobic conditions (N2:CO2, 80:20, vol/vol) at 30°C, as previously described ( 68 ). G. metallireducens GS15 (ATCC 53774) was grown in a Fe(III) citrate (FC) medium ( 69 ) with 20 mM ethanol provided as the sole electron donor and 56 mM Fe(III) citrate as the sole electron acceptor or with 20 mM acetate as the donor and 50 mM Fe(III) oxide as the acceptor. G. sulfurreducens PCA (ATCC 51573) was grown in a medium with 10 mM acetate provided as the sole electron donor and 40 mM fumarate as the sole electron acceptor (NBAF medium) ( 68 ). Co-cultures were initiated with equal amounts of both organisms in anaerobic pressure tubes containing 10 mL of NBF medium (acetate-free NBAF), with 10 mM ethanol provided as the sole electron donor and 40 mM fumarate as the electron acceptor. When noted, additions of anthraquinone-2,6,-disulfonate (AQDS) were made from a concentrated stock to provide a final concentration of 50 µM. In some instances, granular activated carbon [GAC; 8–20 mesh (Sigma-Aldrich)] was added at 0.1 g/10 mL. +Methanosarcina barkeri MS (DSM 800) was obtained from our laboratory culture collection and grown at 30°C in co-culture with G. metallireducens with ethanol as the electron donor, as previously described ( 59 ). Co-cultures were initiated with a 0.5 mL inoculum of both G. metallireudcens and M. barkeri from cultures in the late exponential phase. Analytical techniques Organic acids were monitored with high-performance liquid chromatography, as previously described ( 70 ). Changes in ethanol concentration were monitored with gas chromatography, as previously described ( 4 ). Methane was monitored in the headspace by gas chromatography with a flame ionization detector (SHIMADZU, GC-8A), as previously described ( 71 ). Fe(II) concentrations were determined by first incubating samples for 1 h in 0.5N HCl and then measuring Fe(II) with a ferrozine assay at an absorbance of 562 nm ( 72 ). Illumina sequencing and data analysis For all experimental conditions, total RNA was extracted from triplicate samples at mid-log phase growth when succinate concentrations reached approximately 25 mM using the RNeasy Mini Kit (Qiagen) according to the manufacturer’s instructions. Samples were treated with Turbo DNA-free DNase (Ambion, Austin, TX), and the RNA samples were tested for genomic DNA (gDNA) contamination by PCR amplification of the 16S rRNA gene. mRNA was enriched using the MICROBExpress kit (Ambion), according to the manufacturer’s instructions. Directional libraries were prepared with the ScriptSeq v2 RNA-Seq Library Preparation Kit (Epicentre), and single-end sequencing was performed on a Hi-Seq 2000 platform at the Deep Sequencing Core Facility at the University of Massachusetts Medical School in Worcester, Massachusetts. The program FASTQC ( http://www.bioinformatics.babraham.ac.uk/projects/fastqc/ ) was used to visualize and quality check all raw data. Initial raw non-filtered libraries contained an average of 13175155.5 ± 1,758,892 and 10227370.2 ± 1558219.6 reads in the DIET and quinone-mediated interspecies electron transfer (QUIET) libraries that were ~100 base pairs long. Sequences from all of these libraries were trimmed and filtered with Trimmomatic (bolger 2014) yielding an average of 9,286,241 ± 1665081.9 and 12,393,095 ± 1719373.9 reads for the DIET and QUIET libraries. Mapping of mRNA reads Trimmed and filtered mRNA reads from triplicate samples for the two different co-culture conditions (DIET and QUIET) were mapped against the G. metallireducens strain GS-15 genome ( NC_007517 ) and the G. sulfurreducens strain PCA genome ( NC_002939 ) downloaded from IMG/MER ( img.jgi.doe.gov + ) using ArrayStar software (DNAStar). Common dispersion (Disp) and biological coefficient of variation (BCV) values between DIET and QUIET replicates were calculated with the edgeR package in Bioconductor ( 73 ). Common dispersion and BCV values for DIET and QUIET libraries were Disp = 0.01953 and BCV = 0.1398 and Disp = 0.11752 and BCV = 0.3428, respectively. A multidimensional scaling (MDS) plot was also generated with edgeR software and showed that replicates from the DIET and QUIET libraries clustered together but separately from each other (Fig. S2). Once the quality of RNAseq libraries was determined, differential expression studies were done with the edgeR package in Bioconductor ( 73 ). Genes with P-values <0.05 and fold changes >2 were considered differentially expressed. Using these criteria, 945 G. metallireducens genes and 967 G. sulfurreducens genes were upregulated in DIET-grown co-cultures, and 603 G. metallireducens genes and 848 G. sulfurreducens genes were upregulated in QUIET-grown co-cultures ( Table S2 ). Quantitative RT-PCR Quantitative RT-PCR was conducted with mRNA extracted from triplicate cultures of G. metallireducens wild-type and ∆hcp (Gmet_0280) strains grown by Fe(III) oxide respiration, in co-culture with M. barkeri or in co-culture with G. sulfurreducens. Cells were harvested during the mid-logarithmic phase by centrifugation at 4,000 rpm for 15 min at 4°C. After centrifugation, the pellets were frozen in liquid nitrogen and stored at −80°C until RNA extraction procedures were performed. Total RNA from sample pellets was extracted as previously described ( 74 ). Complementary DNA (cDNA) was generated from mRNA using the Invitrogen SuperScript IV First Strand Synthesis System (ThermoFisher Sci). Primer pairs used for qRT-PCR are provided in Table S3. Three different housekeeping genes were used as external controls; recA that codes for recombinase A, proC that codes for pyrroline-5-carboxylate reductase, and rpoB that codes for the beta subunit of RNA polymerase. Power SYBR green PCR master mix (Applied Biosystems, Foster City, CA) and an ABI 7500 real-time PCR system were used to amplify and quantify all PCR products. Each reaction mixture consisted of forward and reverse primers at a final concentration of 200 nM, 5 ng of gDNA, and 12.5 µL of Power SYBR green PCR master mix (Applied Biosystems). Relative levels of expression of the studied genes were calculated by the 2−ΔΔCT + threshold cycle (CT) method ( 75 ). Mutant construction Primers used for construction of gene replacement mutants and complement strains are listed in Table S4. Deletion mutants were made by replacing the gene of interest with a spectinomycin antibiotic resistance cassette ( 76 ). All restriction digestions were carried out according to the manufacturer’s instructions. PCRs were done using the JumpStart Taq DNA polymerase (Sigma-Aldrich). Primer pairs were used to amplify by PCR flanking regions of approximately 500 bp downstream and upstream of the target genes using the appropriate genomic DNA as a template. PCR products were digested with the AvrII (CCTAGG) (NEB, Beverly, MA) restriction endonuclease, ethanol precipitated, and ligated with T4 DNA ligase (NEB). The ligation reaction was loaded onto a 1% agarose gel, and a 1 kb band was purified using the Qiaquick Gel Extraction Kit (Qiagen) and cloned into a pCR2.1 TOPO cloning vector. Sequences of the cloned products were verified by Sanger sequencing. The spectinomycin cassette was digested with XbaI (TCTAGA) (NEB) from pUC19-Sprr loxP ( 76 ), and the recombinant plasmid was digested with AvrII. The spectinomycin resistance cassette was cloned into the plasmid to complete the construction of the mutant alleles. Plasmids bearing mutant alleles were linearized and concentrated by ethanol precipitation. The linearized plasmids were electroporated as described previously ( 76 ). Antibiotics were added for selection purposes only. Replacement of wild-type alleles by mutant alleles was verified by PCR and Sanger sequencing. Gmet_0280 was complemented in trans by amplifying the gene with its native ribosome binding site (RBS) using G. metallireducens genomic DNA as a template. The resulting PCR product was then digested and cloned under the control of a constitutive lac promoter into pCM66 ( 77 ) and electroporated into the Gmet_0280-deficient strain, as previously described ( 76 ). RESULTS AND DISCUSSION Increased expression of G. metallireducens T6SS genes during DIET Comparing gene expression patterns in defined co-cultures growing via DIET versus co-cultures growing with the exchange of diffusible electron shuttles has proven to be an effective strategy for identifying mechanisms for electron transfer during DIET ( 23 , 24 ). Therefore, co-cultures of G. metallireducens and G. sulfurreducens were grown in a medium with ethanol as the electron donor and fumarate as the electron acceptor as previously described ( 9 , 78 ). The two species must cooperate to share electrons in this medium because only G. metallireducens can utilize ethanol as an electron donor, and only G. sulfurreducens uses fumarate as an electron acceptor ( 9 ). AQDS (50 µM) was added to one set of co-cultures to promote quinone-mediated interspecies electron transfer in which the two species remain free-living, and AQDS serves as an electron shuttle between them ( 78 ). However, in the absence of AQDS, the two species must form a tight physical association for direct electron transfer from G. metallireducens to G. sulfurreducens because G. metallireducens cannot produce H2 or formate as electrons shuttle when metabolizing ethanol ( 22 , 23 ). The initial AQDS-amended and unamended co-cultures were sampled for gene expression analysis after they had reduced ca. 25 mM of the 40 mM fumarate available. DIET-grown G. sulfurreducens had higher transcript abundances for genes previously found to be important in DIET ( Table S2c ). These included genes for over 25 c-type cytochromes, including omcS, omcB, omcX, and omcI, which encode multi-heme, outer-surface c-type cytochromes. The gene coding for the pilin monomer, PilA, which is assembled into electrically conductive pili ( 79 ), was also 63 times more highly expressed in DIET-grown cells. Transcripts for 19 different c-type cytochrome genes were also >twofold more abundant in DIET-grown G. metallireducens ( Table S2a ). These included Gmet_0930, which codes for an octaheme outer membrane c-type cytochrome and Gmet_0910, the gene for the outer membrane c-type cytochrome, OmcF, from the PccF porin-cytochrome complex ( 59 ). Both Gmet_0930 and Gmet_0910 are important for Fe(III) oxide reduction and DIET-based growth ( 17 , 59 , 80 ). Gmet_2029, which codes for a lipopolysaccharide protein likely to be involved in biofilm formation and required for Fe(III) oxide reduction ( 80 ), was expressed more than threefold higher in DIET-grown cells, but pilA was not more highly expressed by DIET-grown G. metallireducens cells ( Table S2a ). Other genes that would not be expected to be involved in electron transfer were also more highly expressed in G. metallireducens cells growing by DIET ( Table 1 ; Table S2a ). The two genes with the greatest increase in abundance of transcripts in DIET- versus QUIET-grown cells were Gmet_2080 and Gmet_2078, annotated as “T6SS needle tube protein TssD” and “T6SS protein ImpB.” Transcripts for other T6SS proteins were also much more abundant in DIET- versus QUIET-grown G. metallireducens ( Table 1 ). This included 13 of the genes needed to construct the T6SS nanomachine in other microbes ( 26 , 31 , 81 ). Furthermore, genes coding for all putative T6SS effectors, immunity proteins, and effector chaperones in G. metallireducens, with the exception of Gmet_0291 which codes for a putative chaperone protein, were at least five times more highly expressed in DIET-grown G. metallireducens cells ( Table 1 ). Only some of the G. sulfurreducens T6SS-related genes were highly expressed in DIET- versus QUIET-grown cells ( Table 1 ). TABLE 1 Differences in transcript abundance for all 13 core T6SS genes in G. metallireducens and G. sulfurreducens grown via DIET compared to QUIET and for genes coding for putative T6SS effector and associated immunity proteins (P-value cutoff ≤0.05) + +a + + Gene Name Annotation Fold-change DIET vs QUIET +P-value +G. metallireducens core T6SS genes Gmet_0273 tssL Membrane core complex protein 129.56 1.38 × 10−6 + Gmet_0274 tssM Membrane core complex protein 47.98 8.99 × 10−26 + Gmet_0275 tssA Baseplate complex protein 91.34 2.71 × 10−23 + Gmet_0278 tssB Sheath protein 703.55 1.71 × 10−21 + Gmet_0279 tssC Sheath protein 44.18 4.29 × 10−27 + Gmet_0280 hcp Inner tube protein 1397.32 1.74 × 10−30 + Gmet_0286 vrgG Puncturing device 44.88 9.12 × 10−15 + Gmet_3310 tssJ Membrane core complex protein 27.08 5.50 × 10−10 + Gmet_3311 tssK Baseplate complex protein 5.69 4.86 × 10−8 + Gmet_3312 tssE Baseplate complex protein 44.53 3.93 × 10−3 + Gmet_3313 tssF Baseplate complex protein 136.45 7.68 × 10−7 + Gmet_3314 tssG Baseplate complex protein 52.36 1.78 × 10−3 + Gmet_3315 tssH ClpV1 protease involved in sheath recycling 11.47 5.50 × 10−11 + +G. metallireducens effector/immunity genes Gmet_0284 tsi1 NTF-domain protein (immunity protein) 14.01 1.02 × 10−8 + Gmet_0285 tse1 Peptidoglycan-binding D-alanyl-D-alanine carboxypeptidase 15.92 9.05 × 10−10 + Gmet_0287 tse2 Fatty acid metabolism protein 5.16 7.36 × 10−5 + Gmet_0288 PAAR-like DUF4150 domain protein 126.78 1.76 × 10−6 + Gmet_0290 tse3 PGAP1 domain protein; phospholipase 7.71 1.31 × 10−8 + Gmet_0291 DUF2169 domain chaperone protein ND +G. sulfurreducens core T6SS genes GSU3165 tssL Membrane core complex protein ND GSU3166 tssM Membrane core complex protein 2.81 0.03 GSU3167 tssA Baseplate complex protein 250.65 3.90 × 10−5 + GSU3172 tssB Sheath protein ND ND GSU3173 tssC Sheath protein 23.07 3.80 × 10−7 + GSU3174 hcp Inner tube protein 11.76 0.0003 GSU3177 vrgG Puncturing device ND ND GSU0428 tssJ Membrane core complex protein ND ND GSU0429 tssK Baseplate complex protein 4.80 0.0003 GSU0430 tssE Baseplate complex protein ND ND GSU0431 tssF Baseplate complex protein ND ND GSU0432 tssG Baseplate complex protein ND ND GSU0433 tssH ClpV1 protease involved in sheath recycling 4.33 0.007 +G. sulfurreducens effector/immunity genes GSU3168 tse1 Fatty acid metabolism protein ND ND GSU3169 PAAR-like DUF4150 domain protein ND ND GSU3171 tse2 PGAP1 domain protein; phospholipase ND ND GSU3176 tse3 lysM domain protein ND ND GSU3181 tse4 Fatty acid metabolism protein 51.42 0.01 GSU3182 PAAR-like DUF4150 domain protein ND + +a + + ND, no difference; tse, T6SS effector protein; tsi, T6SS immunity protein. Disrupting the hcp gene in G. metallireducens accelerates adaption to DIET The high expression of T6SS genes in G. metallireducens during growth via DIET was surprising because a primary function of the T6SS is elimination of competing species ( 31 , 32 , 34 , 82 + – + 84 ). To determine whether expression of the T6SS by G. metallireducens impacted DIET, the gene for the Hcp needle-tube protein (Gmet_0280), the most highly differentially expressed gene in DIET- versus QUIET-grown cells ( Table S2A ), was disrupted by replacing the gene with a spectinomycin resistance cassette. As previously described ( 9 ), co-cultures established in ethanol-fumarate medium with wild-type G. metallireducens required over 25 d to begin DIET, monitored as the accumulation of succinate from fumarate reduction ( Fig. 1A ). In contrast, there was a shorter lag in adaption to DIET in co-cultures initiated with the Hcp-deficient strain of G. metallireducens ( Fig. 1A ). Large aggregates (1–2 mm diameter) were visibly apparent in the co-cultures with the Hcp-deficient strain of G. metallireducens even when the co-cultures were first established. In contrast, as previously reported ( 9 ), in co-cultures established with wild-type G. metallireducens, large aggregates only appeared after multiple successive transfers of the co-cultures. The Hcp-deficient strain also produced visible aggregations in Fe(III) citrate medium, which was not observed in the wild-type strain (Fig. S3). Fig 1 The impact of deletion of a key T6SS component on DIET. Metabolism of (A) DIET co-cultures initiated with the wild-type or hcp-deficient strain of G. metallireducens and wild-type G. sulfurreducens, (B) GAC-supplemented co-cultures initiated with the wild-type or hcp-deficient strain of G. metallireducens and wild-type G. sulfurreducens, (C) QUIET co-cultures initiated with the wild-type or hcp-deficient strain of G. metallireducens and wild-type G. sulfurreducens, and (D) DIET co-cultures initiated with the wild-type or GSU3174-deficient strain of G. sulfurreducens and wild-type G. metallireducens. Each point and error bars represent the average and standard deviation of triplicate measurements. The Hcp-deficient G. metallireducens strain did not have a substantial advantage over wild-type cells if granular activated carbon was added to the co-cultures ( Fig. 1B ). In the presence of GAC, which is electrically conductive, G. metallireducens and G. sulfurreducens attach to the GAC surface rather than producing dual-species aggregates, and the cells are not close enough for DIET via electrically conductive pili or c-type cytochromes ( 60 ). Deletion of the genes for these biological electrical connections does not inhibit DIET in the presence of GAC, suggesting that GAC is a highly effective conduit for DIET ( 60 ). Co-cultures established with the Hcp-deficient G. metallireducens or wild-type G. metallireducens grew at the same rate when AQDS was provided as an electron shuttle for QUIET ( Fig. 1C ). Like GAC, AQDS also eliminates the need for direct cell-to-cell contacts for interspecies electron transfer ( 78 ). Disrupting the gene for Hcp from G. sulfurreducens (ΔGSU3174) did not substantially decrease the lag time required for initiation of co-culture metabolism under conditions that require DIET for growth ( Fig. 1D ). This is consistent with the observation that G. sulfurreducens did not increase expression of genes for most T6SS components in DIET- versus QUIET-grown cells ( Table 1 ). To determine the potential impact of T6SS expression on Geobacter interactions with methanogens, co-cultures of G. metallireducens and M. barkeri were initiated as previously described ( 10 ) in a medium with ethanol as the electron donor. As previously described ( 10 ), there was a lag period of more than 30 d in co-cultures initiated with wild-type G. metallireducens ( Fig. 2 ). In contrast, there was very little lag in co-cultures initiated with the Hcp-deficient G. metallireducens strain. While studies have focused on T6SSs targeting bacterial and eukaryotic cells ( 85 , 86 ), the effect of T6SSs on archaeal cells requires further study ( 87 ). Fig 2 Methane production from DIET co-cultures initiated with the wild-type or hcp-deficient strain of G. metallireducens and wild-type M. barkeri during the first transfer. Each point and error bars represent the average and standard deviation of triplicate measurements. Disrupting the hcp gene in G. metallireducens has an unexpected impact on extracellular electron transfer The impact of the hcp deletion on extracellular electron transfer was evaluated. The Hcp-deficient strain grew faster than wild-type G. metallireducens when insoluble Fe(III) oxide was provided as the electron acceptor ( Fig. 3 ). A complement strain containing the hcp gene in trans grew at rates similar to the wild-type strain, demonstrating that elimination of the T6SS impacted extracellular electron transfer capabilities. Fig 3 Fe(II) production by wild-type G. metallireducens, Δhcp, and complemented Δhcp strains during growth with acetate (20 mM) provided as the electron donor and Fe(III) oxide (50 mM) provided as the electron acceptor. Results and error bars represent triplicate cultures. The growth rate for Δhcp was 1.5 (P-value = 0.002) and 1.4 (P-value = 0.02) times greater than wild-type or complemented Δhcp strains. Quantitative RT-PCR ( Fig. 4 ) revealed that, compared to the wild-type strain, the Hcp-deficient strain of G. metallireducens more highly expressed genes for key outer-surface components previously shown to be important in extracellular electron transfer ( Fig. 4 ; Table S5). These included genes for PilA, the monomer that is assembled into electrically conductive pili that are required for Fe(III) oxide reduction and DIET ( 25 ) as well as genes for the outer surface c-type cytochromes (Gmet_0930 and Gmet_0910) and a lipopolysaccharide protein (Gmet_2029) that are required for Fe(III) oxide reduction and expected to play an important role in DIET ( 23 , 59 , 80 ). Fig 4 Results from quantitative RT-PCR using primers targeting various genes that code for proteins shown to be involved in extracellular electron transfer. mRNA used as a template was extracted from cultures of G. metallireducens growing by Fe(III) respiration with acetate (20 mM) as the electron donor and Fe(III) oxide (50 mM) as the electron acceptor (Fe(III)-oxide), growing in co-culture with M. barkeri (DIET GM-MB) or growing in co-culture with G. sulfurreducens (DIET GM-GS). Results were calculated from triplicate biological and technical replicates using three different housekeeping genes as references (proC, recA, and rpoB). * represents P-values <0.05; ** represents P-values <0.01. Further details regarding qRT-PCR results and P-values are available in Table S5. Increased expression of these genes is a likely explanation for the accelerated Fe(III) oxide reduction and may also have contributed to accelerated DIET. The biosynthetic and energetic costs of deploying T6SS machinery are high ( 88 ). Therefore, it may be that eliminating some of this cost by deleting hcp enabled a greater investment in the expression of outer-surface proteins important for extracellular electron transfer. Notably, T6SS genes were not upregulated in G. metallireducens/G. sulfurreducens co-cultures that had undergone long-term adaptation to growth via DIET ( 23 ), suggesting that lowering expression of T6SS genes is part of G. metallireducens’ adaptive response to DIET-based growth. Implications The results demonstrate that the expression of a T6SS can be detrimental for the establishment of DIET consortia and offer a new insight into the mechanisms by which conductive materials might facilitate DIET. It has previously been considered that conductive materials that are larger than cells, such as GAC, biochar, or carbon cloth accelerate the initiation of DIET because of the following: (i) expression of electrically conductive pili and some outer-surface cytochromes is no longer necessary, conserving energy and (ii) it is easier for an electroactive microbe to establish electrical contact with a large conductive surface than small, disperse electrical contacts on another microbial cell ( 1 , 89 ). The results presented here suggest that another benefit, in some instances, may be that conductive surfaces alleviate the need for close physical contact between DIET partners ( 60 + – + 62 ). Thus, DIET partners can “socially distance” to avoid the possible negative impact of close physical associations as electrons zoom through the conductive material enabling the cells to connect remotely. +G. metallireducens’ high expression of its T6SS is clearly not in its best interest in the context of DIET in a defined laboratory co-culture, but G. metallireducens seems unlikely to exemplify the Geobacter species that participate in DIET in natural communities. G. metallireducens was recovered from an enrichment culture that was selected for microbes rapidly growing via Fe(III) oxide reduction ( 69 , 90 ), conditions likely to favor interspecies competition, not cooperation. Not all Geobacter species possess a T6SS ( Table S1 ). Many other microbes that participate in DIET lack a T6SS, including Prosthecochloris aestaurii ( 16 ), Syntrophus aciditrophicus ( 8 ), Rhodoferax ferrireducens ( 13 ), Desulfovibrio sp. JY ( 15 ), and Rhodopseudomonas palustris ( 18 , 20 ) ( Table S1b ). The finding that deletion of a major gene necessary for T6SS function in G. metallireducens unexpectedly increased the expression of key components for extracellular electron transfer emphasizes a frequent problem in studies of Geobacter electromicrobiology. Adaption to gene deletions often results in changes in the mutant’s physiology beyond the direct function of the missing protein ( 65 , 91 , 92 ). Thus, multiple experimental approaches are warranted when developing models for Geobacter extracellular electron exchange. DATA AVAILABILITY Illumina sequence reads have been submitted to the SRA NCBI database under BioProject PRJNA722959 and Biosample SAMN18796025 . SUPPLEMENTAL MATERIAL The following material is available online at https://doi.org/10.1128/spectrum.00941-23 . 10.1128/spectrum.00941-23.SuF1 Supplemental figures, tables, and text spectrum.00941-23-s0001.pdf Figures S1, S2, and S3; Tables S3, S4, and S5. 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